Inositol Phosphates: Linking Agriculture and the Environment

Inositol Phosphates

LINKING AGRICULTURE AND THE ENVIRONMENT

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Inositol Phosphates

LINKING AGRICULTURE AND THE ENVIRONMENT

Edited by

Benjamin L. Turner

Smithsonian Tropical Research InstituteBalboa, Ancón, Republic of Panama

Alan E. Richardson

CSIRO Plant IndustryCanberra, Australia

and

Edward J. Mullaney

United States Department of AgricultureNew Orleans, USA

CABI is a trading name of CAB International

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v

Contents

Contributors viiPreface ixAcknowledgements xi

1. Nomenclature and Terminology of Inositol Phosphates: Clarification and a Glossary of Terms 1Stephen B. Shears and Benjamin L. Turner

2. Identification of Inositol Phosphates by Nuclear Magnetic ResonanceSpectroscopy: Unravelling Structural Diversity 7Pushpalatha P.N. Murthy

3. High-performance Chromatographic Separations of Inositol Phosphates and Their Detection by Mass Spectrometry 23William T. Cooper, Matthew Heerboth and Vincent J.M. Salters

4. Origins and Biochemical Transformations of Inositol Stereoisomers and Their Phosphorylated Derivatives in Soil 41Michael F. L’Annunziata

5. Isolation and Assessment of Microorganisms That Utilize Phytate 61Jane E. Hill and Alan E. Richardson

6. Phytate-degrading Enzymes: Regulation of Synthesis in Microorganismsand Plants 78Ralf Greiner

7. Phytases: Attributes, Catalytic Mechanisms and Applications 97Edward J. Mullaney and Abul H.J. Ullah

8. Seed Phosphorus and the Development of Low-phytate Crops 111Victor Raboy

9. Phytase and Inositol Phosphates in Animal Nutrition: Dietary Manipulation and Phosphorus Excretion by Animals 133Xin Gen Lei and Jesus M. Porres

vi Contents

10. Environmental Implications of Inositol Phosphates in Animal Manures 150April B. Leytem and Rory O. Maguire

11. Ligand Effects on Inositol Phosphate Solubility and Bioavailabilityin Animal Manures 169Thanh H. Dao

12. Inositol Phosphates in Soil: Amounts, Forms and Significanceof the Phosphorylated Inositol Stereoisomers 186Benjamin L. Turner

13. Abiotic Reactions of Inositol Phosphates in Soil 207Luisella Celi and Elisabetta Barberis

14. Interactions Between Phytases and Soil Constituents: Implicationsfor the Hydrolysis of Inositol Phosphates 221Timothy S. George, Hervé Quiquampoix, Richard J. Simpsonand Alan. E. Richardson

15. Plant Utilization of Inositol Phosphates 242Alan E. Richardson, Timothy S. George, Iver Jakobsen and Richard J. Simpson

16. Inositol Phosphates in Aquatic Systems 261Ian D. McKelvie

Index 279

vii

Contributors

Barberis, Elisabetta, University of Turin, DIVAPRA Chimica Agraria, via Leonardo da Vinci 44, Grugliasco,10095 Torino, Italy

Celi, Luisella, University of Turin, DIVAPRA Chimica Agraria, via Leonardo da Vinci 44, Grugliasco, 10095Torino, Italy

Cooper, William T., Department of Chemistry and Biochemistry, Florida State University, Tallahassee, FL32306, USA

Dao, Thanh H., United States Department of Agriculture–Agricultural Research Service, Beltsville AgriculturalResearch Center, Room 121, 10300 Baltimore Avenue, Building 306 BARC-EAST, Beltsville, MD 20705,USA

George, Timothy S., Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, UK

Greiner, Ralf, Federal Research Centre for Nutrition and Food, Centre for Molecular Biology, Haid-und-Neu-Straße 9, D 76131 Karlsruhe, Germany

Heerboth, Matthew, Department of Chemistry and Biochemistry, Florida State University, Tallahassee, FL32306, USA

Hill, Jane E., Environmental Engineering Program, Yale University, 9 Hillhouse Avenue, PO Box 8286, NewHaven, CT 06520, USA

Jakobsen, Iver, Risø National Laboratory, Biosystems Department, Roskilde, DK 4000, Denmark

L’Annunziata, Michael F., The Montague Group, PO Box 5033, Oceanside, CA 92052, USA

Lei, Xin Gen, Department of Animal Science, Morrison Hall 252, Cornell University, Ithaca, NY 14853, USA

Leytem, April B., United States Department of Agriculture–Agricultural Research Service, Northwest Irrigationand Soils Research Laboratory, 3793 N. 3600 E., Kimberly, ID 83341, USA

Maguire, Rory O., Crop and Soil Environmental Sciences, Virginia Tech, Box 0404, Blacksburg, VA 24061,USA

McKelvie, Ian D., Water Studies Centre and Chemistry Department, School of Chemistry, Monash University,Clayton, Victoria 3800, Australia

viii Contributors

Mullaney, Edward J., United States Department of Agriculture–Agricultural Research Service, Southern RegionalResearch Center, 1100 Robert E. Lee Blvd, New Orleans, LA 70124, USA

Murthy, Pushpalatha P.N., Department of Chemistry, Michigan Technological University, 1400 TownsendDrive, Houghton, MI 49931, USA

Porres, Jesus M., Departamento de Fisiología, Universidad de Granada, Granada, Spain

Quiquampoix, Hervé, Unité de Science du Sol, INRA-ENSAM, 2 Place Pierre Viala, 34060 MontpellierCedex 1, France

Raboy, Victor, United States Department of Agriculture–Agricultural Research Service, Small Grains and PotatoGermplasm Research Unit, 1691 S. 2700 W., Aberdeen, ID 83210, USA

Richardson, Alan E., CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia

Salters, Vincent J.M., National High Magnetic Field Laboratory and Department of Geological Sciences,Florida State University, Tallahassee, FL 32306, USA

Shears, Stephen B., Laboratory of Signal Transduction, National Institute of Environmental Health Sciences,NIH, DHSS, Research Triangle Park, PO Box 12233, NC 27709, USA

Simpson, Richard J., CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia

Turner, Benjamin L., Smithsonian Tropical Research Institute, Apartado 0843-03092, Balboa, Ancón,Republic of Panama

Ullah, Abul H.J., United States Department of Agriculture–Agricultural Research Service, Southern RegionalResearch Center, 1100 Robert E. Lee Blvd, New Orleans, LA 70124, USA

ix

Preface

Inositol phosphates are a group of organic phosphorus compounds found widely in the natural envi-ronment. They are common in eukaryotic organisms, especially plants, where they constitute most ofthe phosphorus in seeds. Soils and aquatic sediments also contain large amounts of inositol phos-phates, some of which occur in forms that have not been detected anywhere else in nature.

The abundance of inositol phosphates in nature means that they are of widespread interest inthe ecological and environmental sciences. However, it is in the science of animal nutrition that inos-itol phosphates have become a topic of considerable interest. This stems from the fact that mono-gastric animals cannot digest phytate (salts of myo-inositol hexakisphosphate), the most abundantinositol phosphate in cereal grains. Supplemental phosphate is therefore required in the diets of pigsand poultry to maintain productivity.

A consequence of phosphate supplementation is that animal manure can contain considerableconcentrations of phosphorus. Not only does this represent a financial loss to the producer, but it alsocontributes to one of the most pervasive forms of environmental pollution from modern agriculture.Long-term application of manure to agricultural land leads to an accumulation of phosphorus in thesoil and a gradual increase in phosphorus transport in runoff to water bodies. Such diffuse pollutionis now widespread and there are numerous examples of regional-scale water quality deterioration inareas of intensive livestock operations. Two well-publicized examples are the Chesapeake Bay, USA,and the Gippsland Lakes, Australia. In both cases the problems have been severe and public – thehigh-profile detection of the neurotoxin-producing dinoflagellate Pfiesteria piscicida in the ChesapeakeBay being a particular cause for concern.

To address this issue, several strategies of dietary manipulation have been developed to improvethe ability of monogastric animals to digest phytate. These include the use of ‘low-phytate’ grains –mutants selected for the low concentration of inositol phosphate in their seed – and the developmentof transgenic animals that produce phytase, an enzyme that degrades phytate but is not naturallypresent in the guts of monogastric animals.

By far the most successful strategy, however, has been the supplementation of animal diets witha microbial phytase. This is now standard practice in most large-scale animal feeding operations andis even mandated by law in some states of the USA. It has proved to be extremely effective in reduc-ing phosphorus excretion in manure and has the added benefit of improving mineral nutrition byreleasing metals from complexation with phytate.

Despite the wealth of information on inositol phosphates in animal nutrition, the environmentalimpacts of manure-derived inositol phosphates and associated dietary manipulations are not wellunderstood. In particular, the fate of the large amount of inositol phosphates being cycled through

agriculture, especially in regions with high animal densities, is unknown. Importantly, only a handfulof studies have assessed the impact of diet manipulation on phosphorus dynamics in the environment.

Inositol phosphates are rapidly and strongly stabilized in soil, which means that they are oftenconsidered to be biologically unavailable and unlikely to be transported in runoff to water bodies.Yet there is growing evidence that inositol phosphates are not as recalcitrant in the environment asonce thought. It is now clear that many terrestrial microorganisms, including those associated withplants, have the capacity to use inositol phosphates. This trait appears widespread, although its eco-logical implications await investigation. Similarly, when inositol phosphates are transported in runoffto water bodies, they can degrade rapidly and contribute to the nutrition of cyanobacteria and otheraquatic organisms linked to eutrophication. Inositol phosphates can therefore no longer be consid-ered ecologically or environmentally benign.

Given the water quality problems associated with intensive livestock production and thewidespread adoption of dietary modifications, there is an urgent need to improve our understandingof inositol phosphates in the environment. This was addressed at a conference held in August 2005in Sun Valley, Idaho, USA, sponsored by the Soil Science Society of America. The meeting, entitled‘Inositol Phosphates in the Soil–plant–animal System: Linking Agriculture and Environment’, wasattended by scientists from a diverse range of disciplines with a common interest in inositol phos-phates. This book is the output from that conference. Written by the invited speakers, it bringstogether critical reviews on the major topics in inositol phosphates in agriculture, ecology and theenvironment. The chapters cover three major themes:

1. State-of-the-art analytical methodology for assessing inositol phosphates in environmental sam-ples, including nuclear magnetic resonance (NMR) spectroscopy and mass spectrometry.2. Inositol phosphates in animal nutrition, including the latest research on plant and microbialphytases, their interactions in soil and the manipulation of animal diets with phytase supplementsand low-phytate grains.3. Inositol phosphates in the environment, including the amounts, forms and behaviour in soilsand aquatic systems, their biological availability and the fate of manure-derived inositol phosphatesin the environment.

By covering all major aspects of inositol phosphates in agriculture and the environment, the book willserve as a unique reference source on this emerging topic. We hope that it will benefit those tryingto unravel the complexity of inositol phosphates in the environment and reveal what is already knownto a wider audience.

The inositol phosphate conference in 2005 was held a quarter of a century after the publicationof Dennis Cosgrove’s seminal text Inositol Phosphates: Their Chemistry, Biochemistry, and Physiology (ElsevierScientific, Amsterdam). Formerly of CSIRO Plant Industry in Canberra, Australia, Cosgrove devotedhis career to understanding inositol phosphates in the environment (an obituary can be found in SoilBiology and Biochemistry, vol. 14, pp. 77–78). His pioneering work in the two decades after he movedwith his family from England to Australia in 1955 laid the foundations for many of the topics in thisvolume. His death in 1981 at the age of 56 marked the end of an era for studies on inositol phos-phates in the environment, but his discoveries remain an inspiration to scientists in this field. We hopethat this volume will go some way towards reinvigorating interest in these fascinating compounds.

Benjamin L. TurnerSmithsonian Tropical Research Institute, Balboa, Ancón,

Republic of Panama

Alan E. RichardsonCSIRO Plant Industry, Canberra, Australia

Edward J. MullaneyUnited States Department of Agriculture, New Orleans, USA

x Preface

xi

Acknowledgements

The meeting would not have been possible without generous support from the Soil Science Societyof America through the Bouyoucos Conference fund. Bouyoucos conferences were established to facil-itate an intense, highly focused examination of a topic of critical importance to soil science. Scientistswith a common interest are brought together in a forum that is not typically possible at large scien-tific meetings, with the aim of establishing personal relationships and promoting the free exchange ofideas. We hope this latest Bouyoucos Conference fulfilled these ideals.

Additional funding was provided by the Agricultural Research Service of the United StatesDepartment of Agriculture through a Professional Activities grant, and the Sun Valley Resort gen-erously provided their conference facilities without charge. We thank those who gave their time topeer-review chapters for this volume – their expertise has contributed to the technical excellence ofits contents. Finally, we thank all the delegates at the inositol phosphate conference for contributingto a vibrant and stimulating few days, and we look forward to the next meeting.

1 Nomenclature and Terminology ofInositol Phosphates: Clarification and a

Glossary of Terms

Stephen B. Shears1 and Benjamin L. Turner2

1Laboratory of Signal Transduction, National Institute of Environmental HealthSciences, NIH, DHSS, Research Triangle Park, PO Box 12233, NC 27709, USA;2Smithsonian Tropical Research Institute, Apartado 0843-03092, Balboa, Ancón,

Republic of Panama

©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment(eds B.L. Turner, A.E. Richardson and E.J. Mullaney) 1

In a book like this one, which brings togetherreviews from scientists working in such diverseareas as analytical chemistry, biochemistry, agron-omy and environmental science, the consolidationof terminology is of considerable importance.This chapter reviews the nomenclature of inositolphosphates and provides a glossary of the termsthat are used throughout this book.

An Overview of Inositol PhosphateNomenclature

Much of what follows is based on the recom-mendations of the International Union of Pureand Applied Chemistry (IUPAC) (IUPAC–IUBCommission on Biochemical Nomenclature(CBN), 1973, 1977; Nomenclature Committee ofthe International Union of Biochemistry, 1989).Previous conferences, most notably the ‘ChiltonConference on Inositol and Phosphoinositides’,held in Dallas, Texas, USA, in 1984, have per-mitted the use of inositol phosphate nomencla-ture that is not IUPAC-approved (Agranoff et al.,1985). The audience at the Chilton conferencewere largely animal biochemists and the numberof known inositol-containing compounds was farsmaller than today. The authors may thereforehave underestimated the potential for confusion

that lay ahead. It is now arguable that theChilton meeting was a missed opportunity toenforce a much-needed, unified nomenclature.

Inositol phosphate terminology continuesto be misused even in the recent literature.For example, ‘phosphoinositide’ is a term thatwas intended to refer only to the inositol lipids(IUPAC–IUB Commission on BiochemicalNomenclature (CBN), 1977). Instead, conceptualdifficulties arise when phosphoinositide is incor-rectly used to describe inositol phosphates (e.g.De Camilli et al., 1996; Luttrell and Lefkowitz,2002; Liu et al., 2004), especially as the physico-chemical properties and biological actions ofthese soluble inositol derivatives are markedlydifferent from those of the membrane-boundinositol lipids.

Unfortunately, even such esteemed bodiesas IUPAC are not immune from error; theirintentions were confounded somewhat when theinositol lipid used to illustrate nomenclature wasnot the naturally occurring D-enantiomer, butthe unnatural L-version (see Agranoff, 1978). Theadoption of a consistent nomenclature clearlycannot eliminate mistakes, but it is an importantfirst step towards limiting their frequency.

The inositol phosphate literature also containsa number of examples of the misuse of chemicalnomenclature, so clarification is appropriate. Forexample, a newcomer to the field would be

forgiven for assuming that ‘IP6’ has only a singlephosphate at the 6-position, when the abbreviationis incorrectly defined as ‘inositol-6-phosphate’ (Leeet al., 2005). The structural relationship betweenthe scyllo- and myo-forms of inositol hexakisphos-phate can be misunderstood when they are erro-neously described as ‘conformers’ (Fisher et al.,2002). Of particular importance to this bookis the term ‘phytate’. This refers to any salt of myo-inositol hexakisphosphate. However, it has some-times been considered that phytate is synonymouswith phytin (Yoshida et al., 1999), even though thelatter term was introduced originally to describecalcium/magnesium phytate, which was thoughtto comprise much of the phytate in seeds (Ashton,1976). Further examples of the continued confu-sion in this field caused by incorrect terminol-ogy are given in a recent review (Michell et al.,2006).

A glossary of terms is provided below as aprelude to this volume. A determined effort hasbeen made to ensure that these terms are consis-tently deployed throughout the various chapters.

Glossary

Conformer. This is one particular spatialarrangement of a molecule in space at any par-ticular moment. For example, two conformers ofmyo-inositol are the so-called ‘chair’ and the‘boat’ arrangements of the ring, the former beingthermodynamically favourable. A switch betweendifferent conformers involves rotation aroundsingle bonds, but no chemical bonds are broken(if bonds were rearranged, and hence the config-uration changed, the two molecules would bestereoisomers). For inositols, the end point ofa conformational change can be a ring-flip,which involves the conversion between two alter-nate chair conformations. This occurs for myo-inositol hexakisphosphate when solution pHincreases past a critical value, whereupon thephosphates switch from being in a 5-equato-rial/1-axial arrangement to a 5-axial/1-equatorialgrouping (see Murthy, Chapter 2, this volume).

Epimer. This is a special case of a pair ofstereoisomers having two or more stereogeniccentres, but differing at only one of these. Forexample, myo-inositol and scyllo-inositolare epimers, because of differences in the spatial

positioning of chemical bonds at one of their sixstereogenic centres.

Epimerization. The process by which twoepimers are interconverted.

Inositol. A cyclitol (cyclohexanehexol) witha hydroxyl group associated with each of the sixcarbon atoms on the ring. See also myo-inositoland scyllo-inositol.

Inositol phosphate. The addition to theinositol ring of an ascending number of phosphategroups gives rise to a series of phosphorylated com-pounds (Table 1.1). The multiplicative prefixes (nopart of which should be italicized) highlight the factthat each carbon atom has only one phosphateattached to it. Thus, ‘bis’, which is Latin in origin,means twice; ‘tris’ is Greek, meaning thrice orthree times; and ‘kis’ is a general prefix from Greekthat means times (Sarma, 2004). This distinguishes‘n’ from ‘n-times’. Thus, if there were an inositolderivative with a chain of three phosphatesattached to a single carbon atom, it would be atriphosphate, not a trisphosphate. The reader whois new to this field may be relieved to know thatinositol triphosphates have not been detected (yet).However, diphosphate groups can be attachedto the inositol ring (Table 1.1). These ‘inositolpyrophosphates’ occur naturally inside cells from awide range of organisms (Shears, 2005). Throughoutthis book the term inositol phosphate is used ina general sense for all phosphorylated inositolspresent in environmental samples.

myo-Inositol. This is one of the nine possi-ble stereoisomers of cyclohexanehexol (Fig. 1.1).In the literature, when the exact nature of thestereoisomer is not defined, it can typically bepresumed to be myo-inositol. In fact ‘Ins’ is an IUPAC-approved term for myo-inositol(Nomenclature Committee of the InternationalUnion of Biochemistry, 1989). The ‘Ins’ abbrevia-tion is not used in this volume, so as not to under-value the significance of the other stereoisomersthat figure prominently in the environment.

myo-Inositol hexakisphosphate. A com-pound in which all six hydroxyl groups of myo-inositol are esterified as phosphates. myo-Inositolhexakisphosphate is a systematic name and is alsopopular in the cell-signalling literature (Irvine andSchell, 2001). Outside that field, this compound ismore usually known as phytic acid. This isstrictly defined as myo-inositol hexakis (dihydrogenphosphate), but the commonly used myo-inositolhexakisphosphate is used in this book.

2 S.B. Shears and B.L. Turner

Nomenclature and Terminology 3

HO HO

HO

HOHO

HO

OHOH

OHOH

OH OH

OH OH

OH

HO

HO

OH

OHOH

HO

HOOH

HO

OH HO

HO

HOHO

OH OH

HOHO

OH

OH OH HO

HO

HO

OH

OH

HO

HO

OH

allo-Inositol

HO

HO

HOOH

OH OH

HO

OH

myo-Inositol

HO

scyllo-Inositol neo-Inositol

L-chiro-(−)-Inositol D-chiro-(+)-Inositol epi-Inositol

HO

muco-Inositol cis-Inositol

Table 1.1. The myo-inositol phosphates and their accepted abbreviations.

Number of IUPAC Common Full name phosphate groups abbreviationa abbreviation

myo-Inositol 0 Ins Insmyo-Inositol monophosphate 1 InsP1

b IP1myo-Inositol bisphosphate 2 InsP2 IP2myo-Inositol trisphosphate 3 InsP3 IP3myo-Inositol tetrakisphosphate 4 InsP4 IP4myo-Inositol pentakisphosphate 5 InsP5 IP5myo-Inositol hexakisphosphate 6 InsP6 IP6Diphospho-myo-inositol

tetrakisphosphate 6 PP-InsP4 PP-IP4Diphospho-myo-inositol

pentakisphosphate 7 PP-InsP5 IP7Bis-diphospho-myo-inositol

tetrakisphosphate 8 [PP ]2-InsP4 IP8

aThe italicization of the P denotes its use as an abbreviation for phosphate, rather than the chemical symbol forphosphorus.bAlthough it is not explicitly stated, we infer that InsP (without a numeric subscript) is actually the IUPAC-preferredabbreviation for myo-inositol monophosphate. However, we recommend InsP1, to avoid confusion with ‘InsP’, which issometimes incorrectly used as a collective abbreviation for inositol phosphates (e.g. Tavares et al., 2002; Woodcocket al., 2003).

Fig. 1.1. The nine stereoisomeric forms of inositol.

Phytase. An enzyme (myo-inositol hexak-isphosphate phosphohydrolase) that initiates thecleavage of one or more phosphate groups frommyo-inositol hexakisphosphate. Several phy-tases are now known to exist and are described indetail in this volume (see Mullaney and Ullah,Chapter 7). Some authors prefer the term phy-tate-degrading enzyme.

Phytate. This refers to any salt of phyticacid. Phytate can be soluble or insoluble and canoccur in both dissolved and precipitated forms.Insoluble phytate involves polyvalent cations (e.g.iron phytate), whereas soluble phytate usuallyinvolves monovalent cations (e.g. sodium phy-tate). However, a recent study concluded that theneutral pentamagnesium salt is the predominantsoluble form in animal cells (Torres et al., 2005).Nevertheless, phytate will precipitate out of solu-tion, in a pH-dependent manner, once a criticalconcentration of divalent cations is exceeded. Inmost cases, myo-inositol hexakisphosphateexists as a salt, in both precipitated and dissolvedforms, and can thus be termed phytate. However,to avoid confusion, the term phytate is used inthis volume only when additional informationabout the cation or solubility is known.

Phytate-degrading enzyme. This is analternative term for phytase that is preferred bysome authors when the in vivo function of theenzyme has not been unambiguously demon-strated (see Greiner, Chapter 6, this volume).

Phytic acid. This is a non-systematic butwidely used alternate name for the free-acid formof myo-inositol hexakisphosphate. As thesalt-free form is unlikely to occur widely in nature,the term myo-inositol hexakisphosphate ispreferred over phytic acid in this volume, althoughthe term phytate is used when the salt is known.Phytic acid should not be used to describe otherphosphorylated stereoisomers such as scyllo-inositol hexakisphosphate.

Phytin. This term was originally intro-duced to describe ‘insoluble’ calcium/magnesiumphytate deposits in the globoids of plant seed(e.g. Ashton, 1976). The term is largely obsolete,because phytate in the seeds of many species isnow known to consist predominantly of magne-sium/potassium salts (Ockenden et al., 2004).The use of ‘insoluble’ as an absolute descriptionof this material also seems unwarranted, as thedeposits are mobilized during seed germination.

Positional isomers. This is a form of struc-tural isomerism in which side chain groups (in this

case phosphates) are found attached to differentcarbons of the inositol ring. That is, atoms arebonded together in a different order, as opposed tostereoisomers, in which the connectivity is thesame. myo-Inositol 1,3,4,5-tetrakisphosphate andmyo-inositol 3,4,5,6-tetrakisphosphate are examplesof positional isomers. The numbering of the car-bon atoms follows rules developed by IUPAC(IUPAC–IUB Commission on Biochemical


HO

HOOH

OHOH

HO

21

3

4

5

6

2

1

6

54

3

( a )

( b )

Fig. 1.2. (a) Agranoff’s turtle and (b) myo-inositol.(From Shears, 2004.) In its most stable chairconformation with 1-axial and 5-equatorialhydroxyl groups, myo-inositol has been said toresemble a turtle (Agranoff, 1978). InternationalUnion of Pure and Applied Chemistry (IUPAC)rules state that the 1-D-numbering of each carbonbegins with the turtle’s front right flipper andproceeds in an anticlockwise direction around thering (viewed from above). The axial hydroxyl istherefore represented by the turtle’s head (positionnumber 2) and the equatorial hydroxyls by thelimbs and tail. For further details of the numberingsystem and stereochemistry of the inositolphosphates the reader is referred to the IUPACrecommendations (IUPAC–IUB Commission onBiochemical Nomenclature (CBN), 1977;Nomenclature Committee of the InternationalUnion of Biochemistry, 1989) and comprehensivereviews published elsewhere (Parthasarathy andEisenberg, 1991; Murthy, 2006).

Nomenclature (CBN), 1973; NomenclatureCommittee of the International Union of Bio-chemistry, 1989). Agranoff’s turtle (see Agranoff,1978; Shears, 2004) provides a timeless, visualmnemonic to the numbering of myo-inositol(Fig. 1.2). Unfortunately, there are no such aids forthe other stereoisomers of inositol.

scyllo-Inositol. This is one of the ninepossible stereoisomers of cyclohexanehexol(inositol). It differs from myo-inositol at onlyone stereogenic centre – i.e. it is an epimer –and is unique in that it has a stable chair con-formation in which all six hydroxyl groups areequatorial to the plane of the ring (Fig. 1.1).

Stereoisomer. This refers to compoundsthat have the same chemical formula, the sameatoms and the same connectivity, but differ inthe fixed spatial positioning of bonds at a partic-ular stereogenic carbon (for the inositols, a stere-ogenic carbon is one lacking a plane ofsymmetry). Hydroxyl groups on the inositolring can be oriented in either an axial or equa-torial manner, which gives nine possible

stereoisomers (Fig. 1.1). These stereoisomers aredistinguished by a configurational prefix, whichmust be italicized (IUPAC–IUB Commission onBiochemical Nomenclature (CBN), 1973). Themost abundant stereoisomer in nature is myo-inositol, but several others occur in plants andanimals. Only four inositol stereoisomers (myo-,neo-, scyllo- and D-chiro-) occur naturally in phos-phorylated forms, predominantly in soils (seeL’Annunziata, Chapter 4, and Turner, Chapter12, this volume).

Turtle. A marine reptile that provides anaide-mémoire for easy recall of the nomenclaturefor numbering the carbon atoms that comprisethe myo-inositol ring (Fig. 1.2; Agranoff, 1978;Shears, 2004).

Acknowledgements

We thank Dr Andrew Riley, University of Bath,UK, and Victor Raboy, USDA–ARS Aberdeen,USA, for their valuable contributions.

Nomenclature and Terminology 5

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Lee, H. J., Lee, S.A. and Choi, H. (2005) Dietary administration of inositol and/or inositol-6-phosphate preventschemically induced rat hepatocarcinogenesis. Asian Pacific Journal of Cancer Prevention 6, 41–47.

Liu, J.W., Anderson, S.N., Meulbroek, J.A., Hwang, S.M., Mukerji, P. and Huang, Y.S. (2004)Polyphosphoinositides suppress the adhesion of Haemophilus influenzae to pharyngeal cells. Lipids in Health andDisease 3 (online-only journal: doi:10.1186/1476-511X-3-20).

Luttrell, L.M. and Lefkowitz, R. J. (2002) The role of beta-arrestins in the termination and transduction of G-pro-tein-coupled receptor signals. Journal of Cell Science 115, 455–465.

Michell, R.H., Heath, V.L., Lemmon, M.A. and Dove, S.K. (2006) Phosphatidylinositol 3,5-bisphosphate: metab-olism and cellular functions. Trends in Biochemical Sciences 31, 52–63.

Murthy, P.P.N. (2006) Structure and nomenclature of inositol phosphates, phosphoinositides, and glycosyl-phatidylinositols. In: Lahiri Majumder, A. and Biswas, B.B. (eds) Biology of Inositols and Phosphoinositides.Springer-Verlag, Berlin, pp. 1–20.

Nomenclature Committee of the International Union of Biochemistry (1989) Numbering of atoms in myo-inositol.Recommendations 1988. Biochemical Journal 258, 1–2.

Ockenden, I., Dorsch, J.A., Reid, M.M., Lin, L., Grant, L.K., Raboy, V. and Lott, J.N.A. (2004) Characterizationof the storage of phosphorus, inositol phosphate and cations in grain tissues of four barley (Hordeum vulgareL.) low phytic acid genotypes. Plant Science 167, 1131–1142.

Parthasarathy, R. and Eisenberg, F. Jr (1991) Biochemistry, stereochemistry, and nomenclature of the inositolphosphates. In: Reitz, A.B. (ed.) Inositol Phosphates and Derivatives. American Chemical Society, Washington,DC, pp. 1–19.

Sarma, N.S. (2004) Etymology as an aid to understanding chemistry concepts. Journal of Chemical Education 81,1437–1439.

Shears, S.B. (2004) How versatile are inositol phosphate kinases? Biochemical Journal 377, 265–280.Shears, S.B. (2005) Telomere maintenance by intracellular signals: new kid on the block? Proceedings of the National

Academy of Sciences of the United States of America 102, 1811–1812.Tavares, P., Martinez-Salgado, C., Ribeiro, C.A., Elono, N., Lopez-Novoa, J.M. and Teixeira, F. (2002)

Cyclosporin effect on rat aorta α1-adrenoceptors and their transduction mechanisms. Journal of CardiovascularPharmacology 40, 181–188.

Torres, J., Domínguez, S., Cerdá, F.M., Obal, G., Mederos, A., Irvine, R.F., Dìaz, A. and Kremer, C. (2005)Solution behaviour of myo-inositol hexakisphosphate in the presence of multivalent cations. Prediction of aneutral pentamagnesium species under cytosolic/nuclear conditions. Journal of Inorganic Biochemistry 99,828–840.

Woodcock, E.A., Mitchell, C. J. and Biden, T. J. (2003) Phospholipase Cδ1 does not mediate Ca2+ responses inneonatal rat cardiomyocytes. FEBS Letters 546, 325–328.

Yoshida, K.T., Wada, T., Koyama, H., Mizobuchi-Fukuoka, R. and Naito, S. (1999) Temporal and spatial pat-terns of accumulation of the transcript of myo-inositol-1-phosphate synthase and phytin-containing particlesduring seed development in rice. Plant Physiology 119, 65–72.


2 Identification of Inositol Phosphatesby Nuclear Magnetic Resonance

Spectroscopy: Unravelling StructuralDiversity

Pushpalatha P.N. MurthyDepartment of Chemistry, Michigan Technological University, 1400 Townsend Drive,

Houghton, MI 49931, USA


At first glance, inositols are deceptively simplemolecules. On closer examination, a host of stereo-chemical, regiochemical, prochiral and conforma-tional issues reveal themselves (Posternak,1965; Parthasarathy and Eisenberg, 1986, 1990).In fact, the International Union of Pure andApplied Chemistry (IUPAC) needed threeattempts and 26 years to agree on a systemof nomenclature that adequately representsthe stereochemical issues involved (IUPACCommission on the Nomenclature of OrganicChemistry and IUPAC–IUB Commission onBiochemical Nomenclature (CBN), 1976; IUBNomenclature Committee, 1989). The complexityis due to the presence of numerous stereochemicalelements in the molecule, including nine stereoiso-mers of the parent inositol moiety (scyllo-, neo-,muco-, etc.), multiple phosphorylated derivatives (63different compounds are possible in the case ofmyo-inositol) and the presence of conformationalisomers (Posternak, 1965; Parthasarathy andEisenberg, 1986, 1990; Murthy, 2006; see Shearsand Turner, Chapter 1, this volume). A completestructural analysis of inositol phosphates thereforerequires that all of these elements be determined.

Structural analysis of inositol phosphatesrequires the extraction of polar, highly chargedmolecules with minimal structural perturbation,chromatographic purification and, finally, estab-lishment of their molecular architecture. As inosi-

tol phosphates can carry numerous negativecharges and often exist in chelated forms, theirextraction and purification poses many challenges.Structures of inositol phosphates can be estab-lished by chemical degradation or nuclear mag-netic resonance (NMR) spectroscopy (reviewed inIrvine, 1986). The procedure for chemical degra-dation involves subjecting a purified and radiola-belled inositol phosphate to a series of chemicaland enzymatic reactions, followed by identifica-tion of the products by co-migration with stan-dards (Fig. 2.1; Grado and Ballou, 1961). This isan elaborate and time-consuming method, whichnormally takes months to complete. In addition, itis an indirect method of structural determinationand the conclusions are therefore ambiguous.

In contrast, NMR spectroscopy is a versatilemethod that can provide complete structuralinformation in a few hours (Derome, 1987;Friebolin, 1993; Claridge, 1999). Some of theadvantages of NMR spectroscopy over conven-tional methods are:

● Inositol phosphates contain three NMR-active nuclei (1H, 31P and 13C), so a detailedpicture of the molecule can be gleaned bycombining information from all nuclei.

● Structural conclusions are direct and unam-biguous and are obtained without the needfor co-migration with standards.

● Information can be obtained about the struc-ture of the parent inositol moiety, the numberand positions of phosphorylation on the ring,and the conformation of the cyclohexane ring.

● Dynamic processes such as conformationalinversion and chemical reactions of inositolphosphates can be investigated.

● Analysis is non-destructive and involvesminimal sample manipulation.

Disadvantages with NMR spectroscopy include:

● Low sensitivity: about 0.1 µmol of the inosi-tol phosphate is required, which correspondsto ~25 µg or 1 ml of ~100 µM sample.

● Inability to distinguish between enan-tiomers: additional experiments with shiftreagents can provide enantiomeric informa-tion, but there are currently no establishedmethods for inositol phosphates.

● The equipment is expensive and the tech-nique is not user-friendly.

NMR spectroscopy has nevertheless been used toobtain a variety of information about inositol phos-phates (Table 2.1). Many of these applications arereviewed here. This chapter is not meant to becomplete or exhaustive, but includes discussion ofthe procedures most likely to be useful for theinvestigation of environmental samples.

Conformational Inversion

Conformational isomers are structural isomersthat are interconverted by rotation around single

bonds (Carey and Sundberg, 2000). The exam-ple of myo-inositol hexakisphosphate is shown inFig. 2.2. The properties of conformational isomers,including size, shape, energy, chemical reactivity,ability to chelate with metal ions and bindinginteractions with proteins, can be markedly dif-ferent. Conformational flexibility of biomoleculeshas a major impact on binding interactions withenzymes and receptors, and therefore on biolog-ical activity. The energy required for rotationaround single bonds is low, with the activationenergy for the chair–chair transition of cyclo-hexane being about 45 kJ/mol (10.8 kcal/mol)(Carey and Sundberg, 2000). This means thatinterconversion between conformational isomersoccurs readily at room temperature and multiplelow-energy conformers exist. Information on pos-sible low-energy conformers at room tempera-ture is therefore necessary to understand bindinginteractions with proteins and metal ions.

NMR spectroscopy has been applied exten-sively to investigate the conformational flexibilityand chair–chair interconversions of a number ofinositol phosphates ( Johnson and Tate, 1969;Costello et al., 1976; Isbrandt and Oertei, 1980;Emsley and Niazi, 1981; Lasztity and Lasztity,1990). In fact, the conformation adopted by myo-inositol hexakisphosphate has been the subject ofmuch debate and was one of the first applica-tions of 31P NMR spectroscopy for structuralinvestigation of inositol phosphates. Johnson andTate (1969) employed 31P NMR spectroscopy toconfirm the structure of myo-inositol hexakispho-sphate and suggested that the conformation wasthe sterically favourable 1-axial/5-equatorial form(Fig. 2.2). However, X-ray crystal data of the

8 P.P.N. Murthy

HO

1. Reduction2. Dephosphorylation

4

5

POO

2

1OH

H H

Periodate5

4OH

OH

OH1

2

Ins(1,4,5)P3

CH2OH

HO

OH

OH

CH2OH

D-Iditol

P

P

P

PP

P

Fig. 2.1. Chemical degradation of myo-inositol 1,4,5-trisphosphate (Ins(1,4,5)P3) to D-iditol.

dodecasodium salt clearly indicated that in solidstate it was in the sterically unfavourable 5-axial/1-equatorial form (Blank et al., 1971).

A number of subsequent NMR studies sug-gested that notwithstanding bulky phosphategroups, conformational inversion to the stericallyhindered form does indeed occur in myo-inositolhexakisphosphate and other inositol phosphates(Isbrandt and Oertei, 1980; Emsley and Niazi,1981; Brigando et al., 1995; Barrientos and

Murthy, 1996; Bauman et al., 1999; Paton et al.,1999; Blum-Held et al., 2001; Volkmann et al.,2002). Numerous theoretical studies also supportthese conclusions (Bauman et al., 1999; Volkmannet al., 2002; Yang et al., 2005). However, the pHdependence of the conformational inversion processand the structural and environmental factorsthat contribute to stabilizing the stericallyhindered form are not completely understood.The fact that the coupling constants in 1H-NMR

Identification by NMR Spectroscopy 9

Table 2.1. Applications of nuclear magnetic resonance (NMR) spectroscopy for the investigation ofinositol phosphates.

Application Representative references

Conformational analysis, including Johnson and Tate (1969), Costello et al. (1976), Isbrandt and activation energy of ring inversion Oertei (1980), Emsley and Niazi (1981), Lasztity and Lasztity of inositol tetrakis-, pentakis- and (1990), Brigando et al. (1995), Barrientos and Murthy (1996), hexakisphosphates Bauman et al. (1999), Paton et al. (1999), Blum-Held et al.

(2001), Volkmann et al. (2002)Structural determination of pure Lemieux et al. (1957, 1958), Brownstein (1959), Dorman et al.

inositol phosphates (1969), Angyal et al. (1974), Angyal and Odier (1982), Cerdanet al. (1986), Lindon et al. (1986), Mayr and Dietrich (1987), Szwergold et al. (1987), Hansen et al. (1989), Barrientos et al.(1994), Johnson et al. (1995), Barrientos and Murthy (1996)

Structural determination of multiple Barrientos et al. (1994), Johnson et al. (1995), Raboy et al.inositol phosphates in a mixture (2000), Dorsch et al. (2003)without separation

Structural determination of impure Volkmann (2002)samples without separation

Acid dissociation constants (pKa) Schmitt et al. (1993), Brigando et al. (1995), Schlewer et al.and protonation sequences of (1999), Blum-Held et al. (2001), Borkovec and Spiess (2004)inositol phosphates at themicroscopic level

Intramolecular hydrogen bonding Felemez and Spiess (2003)in inositol phosphates

Support for theoretical calculations Bauman et al. (1999), Volkmann et al. (2002), Yang et al. (2005)Solid-state NMR spectroscopy Gardiennet et al. (2005)

O−O−

O−

O−

O−

5-axial/1-equatorial1-axial/5-equatorial

O

O−

O−

O−O−

O−O

P

34

−O−O

−O

−O

−O

−O

−O

−O−O

−O

P

O

O

4P

O

O3

12OO

OO P

P

O P−O

O− O−

PO O−

P

OP

OO

O 1 PO

O

O

O

O

P

OP

2O

O

Fig. 2.2. Conformational interconversion of myo-inositol hexakisphosphate.

provide information about the dihedral anglesbetween vicinal protons (coupling constants withvicinal protons) has been particularly useful inthe investigation of chair–chair interconversions(Barrientos and Murthy, 1996). In addition,dynamic NMR spectroscopy has been used toprovide information on the activation energy ofring flipping, while two-dimensional randomdelay exchange spectroscopy (EXSY) has beenapplied to observe the interconversion of mole-cules between different conformations (Fig. 2.3;Bauman et al., 1999; Volkmann et al., 2002).These investigations clearly indicated that myo-inositol hexakisphosphate, scyllo-inositol hexak-isphosphate, all isomers of myo-inositolpentakisphosphate and one isomer of myo-inositoltetrakisphosphate undergo ring–ring interconver-sion from the 1-axial/5-equatorial form to the5-axial/1-equatorial form at room temperature.

Extensive application of 1H, 31P and two-dimensional NMR techniques, supported by the-oretical studies, has indicated that the energy ofconformations is influenced by four factors:

(i) number, substitution pattern and stereochem-istry of phosphate groups on the inositol back-bone; (ii) stereochemistry of the parent inositolring; (iii) physical state (solid or aqueous solution)of the compound; and (iv) properties of the sol-vent, such as pH and counter-ions. From thesestudies, the following generalizations can be made(Barrientos and Murthy, 1996; Bauman et al.,1999; Paton et al., 1999; Blum-Held et al., 2001;Volkmann et al., 2002):

● The pH-dependent conformational prefer-ences of inositol phosphates are unique tothe particular isomer and do not parallelthe behaviour of myo-inositol hexakisphos-phate.

● The presence of four or more equatorialphosphates on the inositol ring induces achange in the conformation from the stericallyunhindered 1-axial/5-equatorial structure tothe sterically hindered 5-axial/1-equatorialconformation at high pH (Fig. 2.4).

● myo-Inositol hexakis- and pentakisphos-phates exist in the 5-axial/1-equatorial con-formation at pH > 10 (Fig. 2.4).

● myo-Inositol 1,4,5,6-tetrakisphosphate, whichcontains four contiguous equatorial phos-phate groups, undergoes conformational in-version to the sterically hindered 5-axial/1equatorial form. However, myo-inositol 1,2,3,4-tetrakisphosphate and myo-inositol 1,2,5,6-tetrakisphosphate do not (Fig. 2.5).

Structural Determination of PurifiedInositol Phosphates

The application of NMR spectroscopy to thestructural determination of inositols dates back tothe early days of NMR spectroscopy in the 1950s(Lemieux et al., 1957, 1958; Brownstein, 1959).Those early investigations quickly revealed thatNMR spectroscopy could distinguish between axialand equatorial groups, because axial protonsoccur at higher field positions compared to equa-torial protons. They also revealed that the chem-ical shifts and coupling constants of protons areinfluenced by the molecular configuration, soNMR techniques held great promise foranalysing the configuration and conformation ofinositol phosphates (Lemieux et al., 1957, 1958;

10 P.P.N. Murthy

4/6

2

4/6 1/3

5

1/3 5 2

4.64.7

4.6

4.5

4.4

4.3

4.2

4.1

4.0

3.9

3.8

3.7

3.6

3.5

3.4

4.5 4.4 4.3 4.2 4.1 4.0 3.9

F1 (ppm)

F2 (ppm)

3.8 3.7 3.6 3.5 3.4

Fig. 2.3. Random delay exchange spectroscopy(EXSY) of myo-inositol hexakisphosphate at pH9.2 with attached 1H spectra. Protons of the 1-axial/5-equatorial conformer are indicated abovethe resonances and protons of the 5-axial/1-equatorial form below. The sweep width in boththe F1 and F2 dimensions was 1355.7 Hz.

Brownstein, 1959). Investigation of the 1H and13C NMR spectra of all nine stereoisomers ofinositol clearly demonstrated that those withthree axial groups (cis-, allo- and muco-) undergochair–chair interconversion at room temperature(Dorman et al., 1969; Angyal et al., 1974; Angyaland Odier, 1982). The interconversion of cis-inositol is slower than the allo- and muco-forms,which interconvert rapidly at room temperature.It was hypothesized that this could be becauseeach hydroxyl group in cis-inositol has to passbetween two hydroxyl groups during the process(Angyal and Odier, 1982).

In the 1980s, NMR spectroscopy (1H, 13Cand 31P) was employed to determine the structureof naturally-occurring inositol phosphates isolatedand purified from cells (Cerdan et al., 1986; Lindonet al., 1986; Mayr and Dietrich, 1987; Szwergoldet al., 1987; Hansen et al., 1989). These studiesestablished the usefulness of NMR spectroscopy forthe structural elucidation of myo-inositol phos-phates, including myo-inositol 1,4,5-trisphosphate,inositol tetrakisphosphates and myo-inositol hexak-isphosphate, and established NMR parameterssuch as chemical shifts, coupling constants andmultiplicity patterns for structural analysis of1H, 13C and 31P NMR spectra of inositol phos-phates. The principal NMR parameters that pro-


21P

PP

P

P

PP

2

1P

PHO

21OH

PP

P

PP

OH

PP

1 2OH

PP

POH

2P

P

P

P

PP

PPP

P2

1P

PP

P

PP

PP

PP

P

PP

PP

PP

PP

21

P1

P

HOP

P

P2

1

Fig. 2.4. Conformational inversion of myo-inositol pentakis- and hexakisphosphates.

PP

PP

P P

P

P

P

P

PP

P

P

PP

PP

PPP

P

P

P

1

11

1 1

1 2

2

2

2

2

2

HO

HO

OHOH

OH

OH

OH

OH

OH

OHOH

OH

Ins(1,2,3,4)P4

Ins(1,2,5,6)P4

Ins(1,4,5,6)P4

Fig. 2.5. Conformational inversion of myo-inositoltetrakisphosphates.

vide structural information include the following(Barrientos et al., 1994; Johnson et al., 1995;Barrientos and Murthy, 1996):

● The number of chemically distinct sets of resonances.The presence of a plane of symmetry wouldresult in four distinct sets of resonances (orfewer if resonances overlap as in myo-inosi-tol hexakisphosphate) and six if there wereno plane of symmetry.

● Chemical shifts of nuclei. These are influencedby the electronic environment, such as theaxial or equatorial orientation of the proton,the nature and number of geminal and vici-nal substituents, and the ionization state.The lack of a phosphate group results in theupfield shift of the α-proton by about0.5–1.0 ppm.

● Multiplicity and coupling constants of resonances.These provide information about the numberand orientation of vicinal protons, dihedralangles and the presence of geminal phos-phates. The splitting pattern of protons onthe inositol ring is due to coupling with twovicinal protons, one on either side (Jax–eqabout 2–3 Hz, Jeq–eq about 2–3 Hz, Jax–axabout 8–10 Hz) and with the phosphorus(JH–P about 8–10 Hz). Therefore, the pres-ence of a phosphate group significantlyaffects the splitting pattern of the inositol ringproton, due to the additional 8–10 Hz1H–31P coupling. Depending on the struc-ture, long-range coupling (W coupling) canalso be detected (Cerdan et al., 1986;Barrientos et al., 1994).

● Two-dimensional NMR experiments. Theseprovide a wealth of information aboutconnectivity and dynamic processes (Derome,1987; Friebolin, 1993; Claridge, 1999). The

application of these general principles for thestructural determination of inositol phosphatesis elaborated in the following section.

Structural Determination of IndividualInositol Phosphates in a Mixturewithout Purification: Application

of Two-dimensional Total CorrelationSpectroscopy

Cells generally contain multiple inositol phos-phates. The metabolism of inositol phosphates istightly interconnected by the action of fast-actingphosphatases and kinases. Therefore, to get acomplete picture of the metabolism of inositolphosphates, changes in the concentration of mul-tiple inositol phosphates must be monitored simul-taneously. Purification of individual inositolphosphates from a mixture is neither easy noralways possible. Analysis of NMR spectra of amixture containing multiple components is diffi-cult due to the inability to assign resonances toindividual molecules, especially in regions wheremultiple resonances overlap. An NMR techniquethat allows the structural assignment of individualcomponents in a mixture without prior separationwould greatly simplify the problem of structuraldetermination.

Two-dimensional total correlation spec-troscopy (TOCSY) experiments can be used todetermine networks of mutually coupled protons(Derome, 1987; Griesinger et al., 1988; Claridge,1999, pp. 201–211). With the addition of a spinlock period the magnetization of H1 is trans-ferred to H2, H3 and H4; i.e. magnetization isrelayed down a chain of contiguous spin-coupledprotons past the vicinal protons (Fig. 2.6). In a

12 P.P.N. Murthy

H1 H2 H3 H4 H5

TOCSY

C C C C C CX

O

P

H

H

H

H H

HHO

HO

OH

OH

OH

Fig. 2.6. Coupling pathway for myo-inositol 5-monophosphate mapped by a TOCSY sequence.

two-dimensional spectrum, H2 will show crosspeaks to H1, H3 and H4. All protons on eachinositol phosphate are part of a connected spinsystem. Therefore, all protons should show con-nectivity either due to direct coupling or long-range magnetization relay. Thus, the TOCSYtechnique provides a way of identifying all theresonances belonging to individual inositol phos-phates ( Johnson et al., 1995). In addition, indi-vidual 1H spectra of each component can beextracted from two-dimensional TOCSY data(Barrientos et al., 1994; Johnson et al., 1995;Raboy et al., 2000; Dorsch et al., 2003). Twoexamples of the use of TOCSY experiments todetermine the structure of individual inositolphosphates in a mixture of inositol phosphatesare described below.

Determination of myo-inositolphosphates in a mixture

The sequential hydrolysis of myo-inositol hexak-isphosphate by phytase produces multiple inositolphosphates. Figure 2.7 shows the TOCSY spec-trum of a mixture of inositol phosphates obtainedby alkaline phytase catalysed hydrolysis of myo-inositol hexakisphosphate. From the one-dimen-sional proton spectrum (top) it is not possible toconfidently deduce either the number or thestructures of inositol phosphates in the mixture.However, the two-dimensional TOCSY spectrumsuggests the presence of three spin systems (i.e.three inositol phosphates) indicated by horizontallines [H], [I] and [ J], with several overlappingresonances. To illustrate the interpretation of a


PP

P

PP

P

P

P

PP

PP

P

PP

1

1

1

2

2

2

HO

HO

HO

F1

(ppm)

3.6

3.8

4.0

4.2

4.4

4.6

4.8

4.8 4.6 4.4 4.2 4.0 3.8 3.6 3.4

2

2

2

4

4,6

4,6

3 1

1,3

1,3,5

6 5

5

F2 (ppm)

[H]

[I]

[J]

[J]

[I]

[H]

Fig. 2.7. Two-dimensional TOCSY spectrum of a mixture of inositol phosphates obtained after 2 h ofalkaline phytase–catalysed hydrolysis of myo-inositol hexakisphosphate (Johnson et al., 1995). Protonspectra are attached. The sweep width in both F1 and F2 dimensions was 1084 Hz. Horizontal lines havebeen drawn to indicate resonances that arise from molecules [H], [I] and [J]. Structures and protonassignments of [H], [I] and [J] are shown.

TOCSY spectrum, Fig. 2.7 is discussed in detailbelow.

One compound gives rise to six peaks asindicated by the arrow [H], a second gives riseto four peaks as indicated by the arrow [I] anda third gives rise to three peaks as indicated bythe arrow [J]. The presence of three sets of res-onances in [J] suggests a plane of symmetry inthe inositol phosphate as well as overlapping res-onances. The relative downfield chemical shiftsof all the 1H resonances suggest that all the car-bons are phosphorylated, so the compound mustbe myo-inositol hexakisphosphate. The chemicalshifts, multiplicity, coupling constants and over-lap of signals from protons in the H-1, H-3 andH-5 positions of the inositol ring provide addi-tional evidence of this assignment. The spectrumof this compound is well documented and hasbeen discussed in detail (e.g. Barrientos andMurthy, 1996).

The presence of four sets of resonances in[I] indicates a symmetrical molecule. The mostnoticeable change in [I] compared to [J] is theupfield shift of one resonance by d ~0.8 ppm tod 3.6 ppm, which indicates that dephosphoryla-tion has occurred on one of the carbons in theplane of symmetry (i.e. either C-2 or C-5). Thepresence of a triplet at d 3.6 ppm rather than aquartet confirms the loss of 1H–31P coupling.The equatorial proton at H-2 has a characteris-tic resonance at ~d 4.8 ppm (a triplet with J ~2–3 Hz), so the upfield shifted of one by ~0.5ppm to d 3.8 ppm resonance. The coupling con-stants of ~8 Hz also provide additional evidencethat the resonance at d 3.6 ppm is due to H-5,because H-2 would give rise to a triplet with J of~2 Hz. Therefore, [I] must be myo-inositol1,2,3,4,6-pentakisphosphate.

The presence of six sets of resonances in [H]indicates a lack of symmetry in the molecule. Themost noticeable change in [H] compared to [I],is the upfield shift of one resonance by about d0.5–3.8 ppm. The triplet splitting pattern ( J ~ 8Hz) indicates the loss of the 1H–31P coupling andsuggests that the resonance must be a proton atH-6 (or H-4) and not H-1 (or H-3), which wouldgive rise to a doublet ( J = 8–9 Hz and 2–3 Hz).Therefore, [H] must be a tetrakisphosphate, eithermyo-inositol 1,2,3,4-tetrakisphosphate or the 1,2,3,6 enantiomer. The connectivity indicated by[H], [I] and [ J] is repeated several times in the

spectrum, both horizontally and vertically,because all the protons of the inositol ring areconnected and therefore show the same connec-tivity pattern. If the composite spectrum is com-plicated with several overlapping resonances, thestructures of the inositol phosphates can be deter-mined by extracting the sub-spectra from the two-dimensional TOCSY data (Johnson et al., 1995).

Inositol phosphates in plant seeds

The TOCSY experiment was employed to inves-tigate the inositol phosphate phenotype of mutantbarley and maize seeds (Raboy et al., 2000;Dorsch et al., 2003). Seeds contain a complex mix-ture of highly phosphorylated inositol phosphates,so the separation and structural determination ofinositol phosphates in such samples pose a formi-dable challenge. The 1H-NMR spectra of inositolphosphates in wild-type barley seeds are shown inFig. 2.8. The one-dimensional 1H-NMR spectrumon top of Fig. 2.8 is complex and contains manyoverlapping resonances. Therefore, it was not pos-sible to confidently deduce either the number orthe structures of inositol phosphates in the mix-ture. The two-dimensional TOCSY spectrum,however, revealed four sets of mutually coupledspin systems (i.e. four inositol phosphates) labelled[11], [12], [14] and [17].

Consideration of the chemical shifts, couplingconstants and multiplicity patterns of each spinsystem, as described above, helped establish thestructures as myo-inositol hexakisphosphate, myo-inositol 1,2,3,4,6-pentakisphosphate, myo-inositol1,2,3,5,6-pentakisphosphate (or its enantiomermyo-inositol 1,2,3,4,5-pentakisphosphate) and myo-inositol 1,2,4,5,6-pentakisphosphate (or its enan-tiomer myo-inositol 2,3,4,5,6-pentakisphosphate)(Dorsch et al., 2003). The assignment of resonancesis indicated in the figure. Thus, the structures ofinositol phosphates in a mixture containing fourhighly phosphorylated derivatives of myo-inositolwere readily ascertained without separation.

In summary, a two-dimensional TOCSYexperiment obviates the need for chromato-graphic separation and provides sufficient infor-mation to unambiguously assign the structures ofclosely related compounds in about 3 or 4 hrather than months.

14 P.P.N. Murthy

Structural Determination of ImpureSamples with Complex Proton

Spectra

When analysing samples that are impure and/ordisplay crowded proton resonances, the TOCSYexperiment may provide spectra that are still toocrowded for unambiguous interpretation. In suchsituations, the presence of the NMR-active het-eroatom 31P (natural abundance 100%) in inosi-tol phosphates can be used as a means to pull out

the 1H spectrum of inositol phosphates. Theproton–proton connectivity information can besorted by the 31P chemical shift attached to thenetwork. Thus, the addition of a TOCSY spinlock mixing period after the heteronuclear multi-ple quantum correlation (HMQC) sequenceallows magnetization transfer on to neighbouringprotons (Fig. 2.9). The result is a 31P-selectedtwo-dimensional TOCSY spectrum; in otherwords, only the proton resonances attached tophosphorus-containing molecules are pulled out


H2

H5

H5

H6

H1H4

H4

H6

H4,H6

H4,H6 H1,H3

H1,H3

H3,H5

H1,H3/H5 [17]

[11]

[14/15]

[12/13]

5.0 4.8 4.6 4.4 4.2 4.0 3.8 3.6 ppm

[17]

[11]

[14/15][12/13]

F2(ppm)

3.6

3.8

4.0

4.2

4.4

4.6

4.8

5.0

5.0 4.8 4.6 4.4 4.2 4.0 3.8 3.6 3.4

F1 (ppm)

5

16

5 3,51,3

4,61,3

6

4

2

4

1,3/5

4,6

2

PP

P

PP

P

P

P

PP

P

P

PP

P

PP

P

P

PP

1

2

[17]

[12]HO

HO

2

2

1

1

OH

[11]

[14]

Fig. 2.8. One-dimensional 1H spectrum (top) and two-dimensional TOCSY spectrum (bottom) of amixture of inositol phosphates extracted from wild-type barley seeds (Dorsch et al., 2003). Protonspectra are attached. Vertical lines have been drawn to indicate resonances that arise from molecules[11], [12], [14] and [17], and the structures and proton assignments of the molecules are shown.

of the complex 1H-NMR spectrum (Friebolin,1993; Braun et al., 1998; Claridge, 1999, pp.241–243).

As an example, the HMQC–TOCSY spec-trum of a mixture of myo-inositol hexakisphosphateand unphosphorylated myo-inositol is shown inFig. 2.10. The resonances downfield of d 4.2 ppm(F2-axis) due to myo-inositol hexakisphosphate are

coupled to phosphorus nuclei (on the F1-axis). Incontrast, the resonances upfield of d 4.0 ppm dueto myo-inositol are not coupled to phosphorusnuclei. Line A indicates that the phosphate at theP-2 position of myo-inositol hexakisphosphate iscoupled to all the protons on the inositol ring, asthese protons are mutually coupled. This is alsotrue for P-1,3, P-4,6 and P-5. Line B shows that

16 P.P.N. Murthy

TOCSYH

H OH

OH

H

OH

H

HHO

H

HO

H1 H2 H3 H4 H5

HMQCC C C C C C

O

P

P

Fig. 2.9. Coupling pathway for myo-inositol 5-monophosphate mapped by a heteronuclear multiplequantum correlation–total correlation spectroscopy (HMQC–TOCSY) sequence.

F2(ppm)

3.2

3.4

3.6

3.8

4.0

4.2

4.4

4.6

4.8

5.0

5.2

5.4

9.0 8.5 8.0 7.5F1 (ppm)

7.0 6.5 6.0

PP

PP

P

P

HH

H HOHO

HO

HO

OH

OH

HHH

1

2

P(5)

P(4,6) P(1,3)

P(2)

A

B

InsP6

H(1

,3) H(5

)

H(4

,6)

H(2

)

H(1

,3,5

)H

(4,6

)

H(2

)

C

Ins

Fig. 2.10. Two-dimensional HMQC–TOCSY spectrum of a mixture of myo-inositol hexakisphosphate andmyo-inositol with attached one-dimensional 31P (top) and 1H (left) spectra (Volkmann, 2002). Vertical andhorizontal lines have been drawn to indicate coupled resonances, and the resonances due to eachcompound are indicated.

the proton at the H-2 position is coupled to allfour 31P resonances, indicating that H-2 is con-nected to a molecule with all four phosphates.This is also the case with H-4,6 and H-1,3,5.The inositol molecule does not contain any 31Pand therefore does not appear in the contourmap; thus the complete proton spectrum of mol-ecules to which the 31P is attached is provided(Volkmann, 2002).

Figure 2.11 shows the spectrum of a mix-ture of myo-inositol hexakisphosphate, myo-inositoland glucose 6-phosphate. The 1H-NMR spec-trum on the left is complicated; it contains manyoverlapping resonances and provides insufficientinformation to make structural assignments. TheHMQC–TOCSY spectrum clearly indicates thepresence of one molecule with one phosphate(line D, glucose 6-phosphate), a second with fourphosphates (line E, myo-inositol hexakisphosphate)and one not coupled to any 31P resonances (myo-inositol). The chemical shifts and the coupling

constants of the 1H resonances and the informa-tion on 31P coupling allow the unambiguous assign-ment of the molecular structures. This techniqueholds great promise for the analysis of compli-cated mixtures of inositol phosphates, as well asin vivo NMR spectroscopy. These applications arecurrently under investigation.

Protonation Sequence at MicroscopicLevel, Acid Dissociation Constants

and Hydrogen Bonding

Potentiometric studies provide acid dissociationconstants (pKa) at a macroscopic level, as wellas overall protonation or dissociation constantsthat describe the molecule as a whole. In inositolphosphates, all phosphates are not equivalent.Therefore, the microscopic pKa values differ andpotentiometric measurements do not provide


F2(ppm)

3.0

3.2

3.4

3.6

3.8

4.0

4.2

4.4

4.6

4.8

5.0

5.2

5.4

InsP6

P(5) P(4,6) P(1,3)

P(2)Glu-6-P

E

D

10.5 10.0 9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0F1 (ppm)

Ins

Fig. 2.11. Two-dimensional HMQC–TOCSY spectrum of a mixture of myo-inositol hexakisphosphate,glucose 6-phosphate and myo-inositol with attached one-dimensional 31P (top) and 1H (left) spectra(Volkmann, 2002). Vertical and horizontal lines have been drawn to indicate coupled resonances, andthe resonances due to each compound are indicated. 1H resonances of glucose 6-phosphate are notindicated because they partially overlap with the myo-inositol resonances as indicated by line D. The 31Presonances of myo-inositol hexakisphosphate and glucose 6-phosphate are indicated on the F1 axis.Lines D and E indicate coupled resonances.

information on the ionization state of individualphosphates. As the chemical shift of 31P is influ-enced mainly by the electronic effects thataccompany protonation and deprotonation, theprotonation sequence at a microscopic level (i.e.the sequence of protonation or deprotonation ofthe various phosphate groups) can be determinedby monitoring the change in the chemical shift of31P as a function of pH. The protonationsequence of myo-inositol hexakisphosphate, myo-inositol 1,4,5-trisphosphate and a number of otherinositol phosphates has been investigated by 31PNMR spectroscopy.

Numerous studies have assessed the pKa valuesof individual phosphates on inositol phosphates(Brigando et al., 1995; Schlewer et al., 1999; Blum-Held et al., 2001; Borkovec and Spiess, 2004). Formyo-inositol hexakisphosphate, Brigando et al. (1995)combined data from potentiometric studies withNMR studies to suggest that the approximatepKa values of the 12 protons are as follows: thefirst proton on the phosphates at the P-2, P-5 andP-1,3 positions are the most acidic, with pKa val-ues less than 2; P-4,6 are less acidic, with pKavalues of 2.6. The pKa values of the second pro-tons of P-5 and P-2 are 6 and 7, respectively,while those of P-1,3 and P-4,6 are ~9, 10, 11and 12, respectively. A similar study was under-taken to determine the microionization constantsof myo-inositol tris- and tetrakisphosphates(Schmitt et al., 1993; Schlewer et al., 1999; Blum-Held et al., 2001; Borkovec and Spiess, 2004),and the sequence of deprotonation of these com-pounds has been elucidated in great detail. Theseinvestigations allow us to pinpoint the exact loca-tion of negative charges at a given pH.

Recently, 1H-NMR methods were employedto investigate hydrogen-bonding interactions ininositol phosphates. Felemez and Spiess (2003)monitored the change in chemical shift of hydroxylprotons as a function of pH and suggested the for-mation of an intramolecular hydrogen bondbetween the 1-hydroxyl and 2-phosphate in myo-inositol 2-monophosphate (Fig. 2.12).

Analysis of Inositol Phosphates inEnvironmental Samples by NuclearMagnetic Resonance Spectroscopy

Although the presence of inositol phosphates interrestrial and aquatic ecosystems has been known

for a long time, little is known about the com-position, cycling, mobility or bioavailability ofinositol phosphates in the environment (reviewedin Turner et al., 2002). The environmental concernsraised by agricultural phosphate contamination,particularly in areas of high livestock density (seeLeytem and Maguire, Chapter 10, this volume),have highlighted the need to accurately monitorinositol phosphates in soils and aquatic sedi-ments. However, the analytical difficulties associ-ated with these studies pose a major challenge.

The extraction and purification of inositolphosphates in soil is complicated by their com-plexation with polyvalent metal ions and associ-ation with complex organic matter such as humicacids (see Celi and Barberis, Chapter 13, this vol-ume). In addition, soils contain uncommon phos-phorylated stereoisomers of inositol (scyllo-, neo- andD-chiro-) in abundance (see Turner, Chapter 12, thisvolume). Methods for the efficient extraction,purification and structural assignment of organicphosphates in soils and other environmental sam-ples have been explored and significant advanceshave been made recently (Turner et al., 2002;Turner and Richardson, 2004; see Cooper et al.,Chapter 3, this volume). The use of one-dimen-sional 31P NMR spectroscopy to analyse alkalinesoil extracts indicated that scyllo-inositol hexak-isphosphate, a compound not reported in plantsand animals to date, is a major component of thesoil organic phosphorus (Turner and Richardson,2004; Turner, Chapter 12, this volume). Theinclusion of a hypobromite oxidation step, whichdestroys all organic phosphates except the inositol

18 P.P.N. Murthy

H

H H

HO

OO

O−

O−

O

O

O O

O

P P

[I] [II]

Fig. 2.12. Intramolecular hydrogen-bondinginteractions in myo-inositol 1,2,6-trisphosphate.[I] and [II] represent alternative intramolecularhydrogen-bonding structures betweenmonoprotonated phosphates and vicinal hydroxylgroups.

phosphates, can significantly help structuralassignment.

Structural identification using one-dimen-sional 31P NMR spectroscopy has several limita-tions, including low sensitivity of 31P compared with1H (6%), narrow spread and poor resolution ofphosphate resonances in the phosphate monoesterregion, and the singlet multiplicity of phosphateresonances that does not provide structural infor-mation about the molecular environment of phos-phates (i.e. other nuclei to which the phosphatesare coupled). The latter is of particular concern,because the presence of rare inositol stereoisomersin soil, such as scyllo-, neo- and D-chiro-inositol,requires structural information about the inositolring connected to phosphates for complete struc-tural identification. Some of these limitations maybe overcome by using two-dimensional techniquessuch as HMQC–TOCSY discussed above. Thismethod may also eliminate the need for hypo-bromite oxidation and the potential structuralchanges associated with it.

For studies involving the concentration andmovement of inositol phosphates in the environ-ment, solid-state NMR would be the preferredmethod of investigation, but the technique pres-ents many problems for application to soils.These include reduced sensitivity and line broad-ening, the presence of paramagnetic ions, narrowspread of 31P resonances so structural informationis hard to obtain and the general difficulty inextracting information from solid-state NMR(Condron et al., 1997). An interesting exampleof the use of solid-state NMR spectroscopy forinositol phosphates was recently described byGardiennet et al. (2005), who used solid-state 31PNMR spectroscopy to detect the presence of themono- and dianionic species of myo-inositol 2-monophosphate.

Summary and Recommendationsfor Future Research

In summary, 1H, 31P and 13C NMR experimentshave revealed many structural details of inositolphosphates, but a number of challenges remain.The intrinsic low sensitivity of NMR spectroscopyand the low endogenous concentrations of inosi-tol phosphates in cells make it difficult to monitorinositol phosphate metabolism in vivo. New tech-niques need to be developed for in vivo studies

and environmental samples. Development ofmethods based on 31P-selected two-dimensionalmethods such as HMQC–TOCSY appears prom-ising for solution NMR. The need to minimizesample manipulation in soils means that the appli-cation of solid-state NMR methods to inositolphosphates in soil samples requires investigation.

Experimental Details

This section provides additional details for theexperiments discussed above. NMR spectra wererecorded on a 400 MHz Varian Unity Inova-400spectrometer. The samples were dissolved indeuterium oxide (0.8 ml) and the pH adjustedto 5.0 with the addition of 1 M NaOH orperdeuterated acetic acid, as necessary. One-dimensional 1H-NMR spectra were obtained at399.943 MHz. 1H chemical shifts were refer-enced to the residual proton absorption of thesolvent deuterium oxide (d 4.67 ppm). For one-dimensional spectra, 16 scans with recycle delayof 6 s between acquisitions were collected. Theacquisition conditions were as follows: spectralwindows 5000 Hz; pulse width 90°. Typically,16–32 scans were collected with recycle delays of4–6 s between acquisitions. The residual waterresonance was suppressed by a 2 s selective pre-saturation pulse.

Two-dimensional EXSY was employed at3°C and pH 10.7. The pulse sequence was atwo-dimensional nuclear Overhauser effect spec-troscopy (NOESY) with a missing time variationincrement of 0.1 s (Claridge, 1999, pp. 326–328).A total of 128t1 increments were obtained, eachconsisting of four transients with a relaxationdelay of 4 s between successive transients. Ashifted Gaussian window was applied in bothdimensions. The data matrix was expanded to a1024 × 1024 real matrix.

TOCSY data-sets were obtained with a 1Hprobe using the pulse sequence of Griesingeret al. (1988). Typically, 128t1 increments werecollected, each consisting of 16–24 transients witha relaxation delay of 6 s between successive tran-sients, using a TOCSY mixing time of 80 ms (asdetermined by one-dimensional TOCSY). Theresidual water signal was suppressed as in theone-dimensional experiment. A Laurentz–Gausswindow was applied in both dimensions, and thedata matrix was expanded to a 1024 × 512 real


matrix. Digital resolutions in the F1 and F2dimensions were ~4 and 2 Hz/point, respectively.

HMQC–TOCSY parameters employed forthe experiments were from Varian Instruments,‘hmqctocsy’ (Varian, 1998). Typically, theparameters were as follows: the F1 dimension was1448.9 Hz and F2 was 1600 Hz. A total of 270 t1increments of 16 transients each were collectedwith a mixing time of 80 ms (as determined byone-dimensional TOCSY) and a relaxation delay

of 4 s. Residual water was suppressed as in theone-dimensional experiment with pre-saturation.

Acknowledgements

The author thanks the National ScienceFoundation (Grant No. CHE-9512445) andMichigan Technological University for funds topurchase a 400 MHz NMR spectrometer.

20 P.P.N. Murthy

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22 P.P.N. Murthy

3 High-performance ChromatographicSeparations of Inositol Phosphates

and Their Detection by Mass Spectrometry

William T. Cooper1, Matthew Heerboth1 and Vincent J.M. Salters2

1Department of Chemistry and Biochemistry, Florida State University, Tallahassee,FL 32306, USA; 2National High Magnetic Field Laboratory and Department of

Geological Sciences, Florida State University, Tallahassee, FL 32306, USA


Mass spectrometry (MS) would appear to be anattractive approach for measuring inositol phos-phates in soils, waters and plant tissues. Thesecompounds are difficult to detect by conventionalultraviolet–visible spectroscopy because they con-tain no chromophoric groups. The commonapproach to quantitatively measuring inositolphosphates is their isolation, oxidation and colori-metric detection. Although this approach is rela-tively simple and straightforward, it can be tedious,time-consuming and subject to interferences. It isalso non-specific, in that all inositol phosphatesgenerate the same signal and they cannot be dis-tinguished by detection alone.

We have demonstrated that electrosprayionization (ESI) combined with high- and ultra-high-resolution MS can be used for qualitativeorganic phosphorus speciation (Llewelyn et al.,2002), i.e. identification of molecular masses andmolecular formulas of individual organic phos-phorus compounds. We also observed that thedetection of individual organic phosphorus com-pounds within a complicated background matrixof natural organic matter could be difficult. Toovercome this problem we recently turned ourattention to coupling liquid chromatographyseparations with inductively coupled plasma(ICP) and ESI–MS, focusing on the qualitativeand quantitative detection of individual inositolphosphates. High-performance size-exclusion

chromatography (HP-SEC) and ion-pairingreversed-phase liquid chromatography (HP-SEC)and reversed-phase high performance liquidchromatography (RP-HPLC) have been evalu-ated for their ability to separate inositol phos-phates based on their degree of phosphorylationand isomeric form.

Unfortunately, none of these techniques canbe fully optimized for inositol phosphate separa-tions because of the limitations imposed by theICP and ESI steps required for MS detection.These ionization techniques require low concen-trations of organic modifiers and volatile salts inthe spray matrices, greatly limiting the separationpotential of the liquid chromatography methods.To date, HP-SEC has proven to be the most ver-satile, though least efficient, separation method.However, we also observed that manipulation ofvarious parameters within the ESI source couldyield resolvable signals for all six inositol phos-phates, allowing detection even when the chro-matographic separation is not entirely satisfactory.

HP-SEC chromatography of inositol phos-phates with ESI–time-of-flight (TOF)–MS detec-tion will be the primary focus of this chapter. ICPionization combined with high-resolution elemen-tal MS provides very sensitive phosphorus-specificdetection and is ideal for quantifying low levelsof organic phosphorus in complex matrices.However, the ESI process, in contrast to ICP

ionization, largely preserves the molecularintegrity of organic phosphates. When combinedwith a high-resolution mass spectrometer, it isuseful for verifying the presence of target analytesand identifying new compounds. For this workwe have combined ESI with a TOF mass spec-trometer that employs reflectron geometry. Thisnew generation TOF instrument includes themost important features of a mass spectrometerused as a chromatographic detector: speed, highmass resolution and good mass accuracy.

Analytical Separations of InositolPhosphates

Note that in general we will restrict this discussionto analytical separations, which include those tech-niques that are primarily designed to maximizethe resolution of individual compounds withincomplex mixtures and quantify them. This is incontrast to preparative separations, which are designedto maximize the recovery and/or purification of atarget compound. Today, analytical separationsare almost always carried out using gas or liquidchromatography, or the very powerful capillaryelectrophoresis technique. Even though gas chro-matography with open-tubular capillary columnsprovides the greatest separation capabilities of allthese methods, the need to volatilize compoundsand move them in the gas phase through a liquid-coated capillary column precludes the routine useof gas chromatography for large, ionic, non-volatile organic phosphate compounds. High-per-formance liquid chromatography (HPLC) andcapillary electrophoresis are thus the techniques ofchoice for separating inositol phosphates.

Separating positional isomers of inositolphosphates is challenging due to the identicalnumber of phosphate groups present. In additionto optimizing the actual separation, a suitabledetection procedure must be selected. Inositolphosphates have no distinct chromophoric groupsand absorb ultraviolet and visible light onlyweakly. Thus, traditional ultraviolet and visibledetectors used in liquid chromatography and cap-illary electrophoresis cannot be used without apost-column derivatization step. Sophisticateddetectors will thus be necessary for widespreadanalytical separations of inositol phosphates,including those involving MS.

We begin with a brief review of classicalmethods used for separating inositol phosphatesbefore discussing more modern, high-resolutiontechniques. The classical methods, although mod-est in terms of capabilities relative to currentlyavailable techniques, nevertheless provided muchof the early information on inositol structures andoccurrence in plants, soils and natural waters.Discussion of these early techniques in relation tothe analysis of environmental samples can befound in a recent review (Turner et al., 2002).

Classical methods

Most of the first truly quantitative separationsof inositol phosphates used low-pressure anionexchange chromatography. Cosgrove (1963,1966, 1969) reported procedures by which a fewinositol phosphate species could be separated.These invariably required harsh solvent condi-tions (e.g. 0.50 M HCl) and resolved only afew target inositols. McKercher and Anderson(1968a,b) also demonstrated the utility of anionexchange columns for inositol phosphate separa-tions. However, inositol phosphates cannot bereliably quantified by ion exchange alone, due tothe co-elution of other organic phosphates. Thiscan be overcome by using hypobromite oxidationprior to chromatography to oxidize interferingsubstances without degrading the higher-orderinositol phosphates (Irving and Cosgrove, 1981).

Paper chromatography (Cosgrove, 1980) andgas–liquid chromatography (Irving and Cosgrove,1982) have also been used to investigate inositolphosphates in environmental samples. The lattertechnique requires derivatization of the polar, ion-ized phosphate groups, a step which adds com-plexity and compromises analytical precision.Paper chromatography can yield fine resolution,but quantification is difficult. However, the tech-nique proved useful for preparative separationsthat preceded a series of MS (L’Annunziata andFuller, 1976), infra-red spectroscopy (L’Annunziataet al., 1977) and proton nuclear magnetic reso-nance (NMR) spectroscopy (L’Annunziata andFuller, 1971) experiments on inositol phosphatesextracted from leaf litter and forest soils. Themicrobially mediated epimerization of radioactive14C(U)-myo-inositol phosphate was even followedby direct autoradiography of paper chromatogra-

24 W.T. Cooper et al.

phy bands (L’Annunziata et al., 1977). Theseexperiments, which were highly sophisticated atthe time, are summarized in detail elsewhere(L’Annunziata, Chapter 4, this volume).

One separation scheme of particular interestused cross-linked dextran gels to separate organicand inorganic phosphorus using Sephadex G-25,with further separation of the organic phosphorusfraction using Sephadex G-50 (Steward and Tate,1971). Although individual organic phosphoruscompounds such as inositols could not be resolved,these latter experiments provided an estimate ofthe relative size of organic phosphorus pools afterhydrolysis with HCl.

Reversed-phase high-performance liquidchromatography

The most straightforward liquid chromatographytechnique is reversed-phase high-performance liq-uid chromatography (RP-HPLC). Note that highperformance refers to liquid chromatographycolumns packed with small (<10 µm diameter) sil-ica gel particles that normally have an activeorganic liquid chemically bonded to their surface.When the liquid is hydrophobic (e.g. octane (C8)or octadecane (C18)), the column is used with apolar mobile phase to produce reversed-phaseseparations based on solute hydrophobicity. Themore hydrophobic the solute, the longer is itsretention time. RP-HPLC separation usuallyrequires a simple mobile phase comprising waterand a polar organic solvent such as methanol,which means that they are ideal when sensitivedetectors or post-column derivatization reactionsare used.

Separation of inositol phosphates based onthe degree of phosphorylation using RP-HPLC isa relatively useful technique, as adding –PO3groups changes the hydrophobicity of the inositolphosphates dramatically. Harland et al. (2004)quantified myo-inositol hexakisphosphate in snackfoods with a C18 column and photodiode arraymultiple wavelength detector after post-columnderivatization with Wade’s reagent. Kasim andEdwards (1998) developed an RP-HPLC systemthat separated myo-inositol trisphosphate throughmyo-inositol hexakisphosphate, as well as posi-tional isomers of the tetra- and pentakisphos-phates, in animal feeds. Casals et al. (2002)

quantified higher-order inositol phosphates usinga similar system that included post-columnderivatization with yttrium, which reduced theabsorbance of an organic–yttrium complex in thepresence of inositol phosphates. Figure 3.1a is aschematic representation of their post-columnreactor and is typical of such reactors, which aredesigned with small volumes to prevent degrada-tion of chromatographic efficiency. Figure 3.1bis a representative chromatogram demonstratingthat straightforward RP-HPLC can, when effec-tively optimized, separate multiple inositol phos-phates, including positional isomers. Note,however, the rather lengthy time required tocomplete the chromatogram.

Ion-pairing reversed-phase high-performance liquid chromatography

Strongly acidic buffers were required in both theRP-HPLC separations of inositol phosphatesdescribed above because inositol phosphates inanionic form are hydrophilic and do not partitioninto the non-polar stationary phase (C18 in thereferences cited) to any extent. Buffers are thusused to suppress ionization and increase the affin-ity of inositol phosphates for the non-polar sta-tionary phase. An alternative to this ionizationsuppression approach is to put hydrophobiccations that form ion pairs with phosphate in themobile phase. By adding such an ion-pairingreagent (e.g. tetrabutylammonium chloride) to themobile phase, the normally hydrophobic station-ary phase becomes coated with the relativelyhydrophobic cation (tetrabutylammonium).Phosphate anions can then form ion pairs withthe exposed positive charges on the stationaryphase surface, providing an ion-exchange-typemechanism. This technique is referred to as ion-pairing RP-HPLC (e.g. Brando et al., 1990). Thepotential of such separations coupled to MSdetection is demonstrated in Fig. 3.2a, in which amixture of organic phosphate standards wasresolved in <20 min (Cooper et al., 2005a). Thissystem was used to qualitatively identify organicphosphorus compounds in a treatment wetland(Fig. 3.2b). This approach holds great promise foranalysis of inositol phosphates because the ion-exchange mechanism appears to provide the bestseparations of these compounds.

HPC Separations and MS Detection 25

Ion exchange and ion chromatography

Currently, ion-exchange chromatography withstrong anion exchange columns is the most suit-able separation technique for inositol phosphates.The ion-exchange mechanism relies on a specificanion (phosphate)–cation (column) interaction.Increasing the number of these interactions byadding phosphate groups increases retentiondramatically. Further, the strength of a phos-phate–cationic site interaction includes a signifi-cant steric component, and thus ion exchangehas proven effective in separating inositol phos-phates and their positional isomers. Skoglundet al. (1998) separated 25 distinct inositol phos-phates using two strong anion exchange columnsfollowed by post-column derivatization and ultra-violet detection.

Ion chromatography is even more powerfulfor inositol phosphate separations because in prin-ciple it allows sensitive conductivity detection. Thetechnique combines an ion-exchange separationcolumn (anion or cation) with a second suppressorcolumn, which reduces the background conduc-

tivity of the mobile phase and allows conductivitydetection of any analyte ion. Ion chromatographythus satisfies both requirements for analytical sep-arations of inositol phosphates: selective retentionmechanisms that allow chromatographic separa-tion of virtually all target analytes, plus sensitivedetection without post-column derivatization.

Phillippy and Johnston (1985) were the first toapply ion chromatography to inositol phosphateanalysis, measuring myo-inositol hexakisphosphatein foodstuffs, although they found post-columnderivatization and ultraviolet detection moresensitive than conductivity. A number of researchgroups subsequently used ion chromatography todetect inositol phosphates. Figure 3.3 demonstratesthe application of such a system for studying theenzymatic dephosphorylation of myo-inositol hexa-kisphosphate, in which 27 inositol phosphates wereseparated and detected (Chen and Li, 2003). Thisparticular separation included a gradient elutionthat allowed weakly retained inositol phosphates tobe separated in a weak mobile phase, and thenmore strongly retained inositol phosphates to beeluted by the increasingly stronger mobile phase.


(a) (b)

Output

Standardconnector

Input PARsolution

StirrerMagnetic

bar

Input eluent

20

0

20

40

mV

60

80

100

30 40

Elution time (min)

50

Inositol polyphosphatesdetected in three pigeonerythrocyte samplesMean � SD(µmol/ml erythrocyte)Ins(1,3,4,6)P4 <0.005Ins(3,4,5,6)P4 0.009�0.005Ins(1,3,4,5,6)P5 3.190�0.34InsP6 0.052�0.019

Ins(

1,3,

4,6)

P4

Ins(

3,4,

5,6)

P4

Ins(

1,3,

4,5,

6)P 5

InsP

6

Fig. 3.1. (a) Post-column reactor for detecting non-absorbing inositols with metal dye reaction.(b) Reversed-phase high-performance liquid chromatography (RP-HPLC) separation of minor inositolphosphates in blood cells. (From Casals et al., 2002. Reprinted with permission from Elsevier Science.)

The chromatogram in Fig. 3.3 demonstrates boththe advantages and disadvantages of gradient elu-tion liquid chromatography, because although the27 inositol phosphates were well resolved in <50min, significant drift in the detector baseline signaloccurred as the composition of the mobile phasechanged.

Capillary electrophoresis

Any review of separations of inositol phosphatesmust include a discussion of capillary elec-trophoresis, a very powerful analytical separationtechnique. It is not a chromatographic technique,because it separates solutes based on elec-trophoretic mobility rather than affinity for a sta-tionary phase. Sample mixtures are loaded into aglass capillary tube filled with electrolyte solution,and a voltage is then applied between the injec-tor and detector ends of the tube. Modern capil-lary electrophoresis evolved from classicalelectrophoresis in the early 1980s when it wasrecognized that the use of small-diameter glasstubes (i.e. capillary tubes of 0.25–0.50 mm i.d.)provided efficient heat transfer and minimizedthe effects of Joule heating. This meant that highvoltages (10–30 kV) could be applied over fairlylengthy distances, resulting in efficient separationof solutes with differing electrophoretic mobility.

Capillary electrophoresis separation of inosi-tol phosphates appears to be straightforward, asaddition of –PO3 groups to the inositol ring shouldchange electrophoretic mobility dramatically. Inaddition, resolution of positional isomers can oftenbe accomplished by adding certain solvents to theelectrolyte solution (e.g. cyclodextrins). However,only a few reports have appeared in the literaturedescribing capillary electrophoresis separations ofinositol phosphates. Henshall et al. (1992) sepa-rated myo-inositol 1,4-bisphosphate, myo-inositol 1-monophosphate, and a mixture of myo-inositol1,4,5-trisphosphate and myo-inositol hexakisphos-phate. They were also able to separate DL-myo-inositol 1-monophosphate and DL-myo-inositol2-monophosphate with a second electrolyte solu-tion. Buscher et al. (1994) used ‘indirect detection’to identify all six inositol phosphate esters by cap-illary electrophoresis, although the resolution ofmyo-inositol penta- and hexakisphosphates waspoor (Fig. 3.4). Taguchi et al. (2000) took a differ-ent approach, using capillary electrophoresis toseparate phospholipids into broad classes, followedby ESI–MS detection. Rather than physically sep-arating the components of each class, they reliedon in-source fragmentation to generate mass spec-tral patterns that could be used to identify severalcomponents within each class.

The principal drawback to capillary elec-trophoresis as a separation technique for inositol


42

8

6

5

3

1

7

0

20,000

40,000

60,000

80,000

100,000

120,000

0 5 10 15 20 25 30Time (min)

CP

S (

P)

(a)

0

2,000

4,000

6,000

8,000

10,000

12,000

0 5 10 15 20 25 30 35 40Time (min)

CP

S (

P)

o-P

o-P23.4 36.1

0

1,000

2,000

3,000

4,000

5,000

0 5 10 15 20 25 30 35 40Time (min)

CP

S (

P)

(b)

1− o-P2− AMP3,4− CP5− ADP6,7− PEP8− ATP

Fig. 3.2. (a) Optimized separation of organicphosphate standards by ion-pair chromatographywith direct, on-line phosphorus-specific inductivelycoupled plasma mass spectrometry (ICP–MS)detection. AMP, ADP, ATP = adenosine mono-, di-and triphosphate; CP = creatine phosphate; PEP= phosphoenolpyruvate. The peak at ~5 min isphosphate. (b) Ion-pair high-performance liquidchromatography (ion-pair HPLC) separation oforganic phosphorus compounds with phosphorus-specific ICP–MS detection from the treatmentwetland inflow (top) and outflow (bottom). CPS =counts per second. (Reprinted from Cooper et al.,2005b.)

phosphates is the low ultraviolet and visible lightabsorbances of the target compounds. Post-col-umn derivatization, which is often used in HPLCseparations, is not possible in capillary elec-trophoresis, so detection normally requires a‘bulk solution’ effect such as decrease in absorp-

tion of a chromophore added to the electrolytesolution. From our experience, this complicatesoptimization of experimental parameters such asapplied voltage, injection time and electrolytecomposition, and even optimized capillaryelectrophoresis systems are far from robust.


0

−0.1

0

0.1

4.9 5.4 5.9

56

432

1

0.06

0.01

0.2

0.3

0.4

0.5

10.0 20.0 30.0 40.0

14.513.512.50.07

0.10

0.13

0.1614 15

1617

1819 20

2122

23

24 2526

27

11

10

12 139

87

15.5 16.5

50.0

Time (min)

CP

S

Fig. 3.3. Separation of 27 inositol phosphate standards by ion chromatography. (From Chen and Li,2003. Reprinted with permission from Elsevier Science.) See citation for identification of inositolphosphates and chromatographic conditions. CPS = counts per second.

0.00−0.02

−0.01

UV

abs

orba

nce

0

0.01

0.02

1.00 2.00

12

3

4

5

6

3.00

Time (min)

4.00 5.00 6.00

Fig. 3.4. Capillary-zone electrophoresis separation of inositol phosphates with indirect ultravioletdetection at 214 nm. (From Buscher et al., 1995. Reprinted with permission from Elsevier Science.) 1 = IP6; 2 = IP5; 3 = IP4; 4 = IP3; 5 = IP2; 6 = IP1.

Furthermore, changes in the electrophoreticmobility of inositol phosphates with the additionof phosphate groups appear to be more compli-cated than first thought.

Mass Spectrometry Detection

There are surprisingly few studies in the litera-ture describing the use of MS for organic phos-phorus detection in natural systems, and evenfewer describing the measurement of inositolphosphates. Several studies attempted to detectlow levels of inositol phosphates in biochemicalmatrices such as red blood cells (Sherman et al.,1986; Hsu et al., 1990, Buscher et al., 1995), butwere largely unsuccessful until the introductionof ESI (Fenn et al., 1989). Organic phosphorus isdominated by the PO4 group(s) that impart(s)polarity and mass. It is thus difficult to ionizeorganic phosphorus molecules without substan-tial fragmentation, complicating the qualitativeidentification of individual compounds. It wasnot until the advent of soft ionization techniques,primarily ESI and matrix-assisted laser desorp-tion ionization, that MS of molecules such asproteins, peptides and inositol phosphatesbecame possible. Kerwin et al. (1994) describedthe role of glycerolphosphoinositols in cell mem-branes using both positive and negative ion-mode ESI–MS. However, it is now apparent thatthe negative ion mode is superior (Hsu and Turk,2000).

Electrospray ionization

The importance of the ionization step in MSdetection of inositol phosphates, as well as thelimitations it imposes on the chemical nature ofthe solvent in which inositol phosphates are dis-solved, means that a brief discussion of the mostimportant ESI process for organic phosphorusMS is necessary. Figure 3.5a is a schematic rep-resentation of an original ESI source design.A liquid solution containing the analyte(s) ispumped through a steel capillary tube held at apotential of 1–3 kV relative to a counter elec-trode in the source. The electrostatic field gen-erated by the applied potential disperses theemerging solution into a fine mist of chargeddroplets. These droplets are rapidly desolvated,leaving charged analytes that are then intro-duced into the mass spectrometer through a‘skimmer cone’, which has the same charge asthe gas-phase ions. The voltage of this cone isimportant, as it needs to be high enough tofocus and guide sufficient ions into the massanalyser to produce a signal, but at the sametime low enough so that it does not induce ‘in-source fragmentation’. Both positive and nega-tive ions can be measured in this way,depending on the polarity of the capillary tipand cone. ESI mass spectra are normally dom-inated by intact molecular ions, often with mul-tiple charges.

Solvent composition is critical for efficientionization by the electrospray method. The


Electrostaticlenses

Capillary

Skimmer

Dryinggas

Firstpumping

stage

Secondpumping

stage

Desolvatinggas

Desolvating chamber

Orifice 2

Ion guide

Ring lens

Orifice 1

(b)(a)

Nebulizer gas

Liquidchromatography

eluent

Cylindricalelectrode

NeedleLiquidsample

Quadrupolemass

spectrometer

Fig. 3.5. Schematic representation of electrospray ionization (ESI) sources: (a) The original design ofFenn et al. (1989); (b) orthogonal design used in the JEOL AccuTOF mass spectrometer. (Courtesy ofJEOL, Inc., Tokyo, Japan.)

solvent must support and encourage ionizationof the analyte(s), but it cannot be better thanthe analyte in taking up the charge appliedby the applied field. Thus, when forming nega-tive analyte ions (e.g. phosphate ions), veryweak bases must be used for buffering. Forexample, we typically use ammonium bicarbon-ate, which is both a weak buffer and quitevolatile. Small amounts of methanol or acetoni-trile are also commonly added to buffered aque-ous solutions to reduce the surface tension ofdroplets.

These constraints make the mobile phasesused in strong anion exchange chromatographyunacceptable for ESI–MS detection, and havebeen one driving force behind the interest infinding alternative separation schemes for inosi-tol phosphates. Dragani et al. (2004) demon-strated a promising approach for analysingglycerophosphoinositol using a β-cyclodextrinbonded phase RP-HPLC system with ESI–MSdetection.

Time-of-flight mass analysis

MS produces information about the masses andabundances of gas-phase ions. It is a very pow-erful analytical technique for determining theelemental composition and chemical formulas ofmolecules, as molecular mass is the single mostimportant piece of information necessary forcharacterizing chemical structures. MS is also animportant quantitative tool because mass spec-trometers are in general very sensitive and willrespond to most compounds in a predictable wayover large concentration ranges. It should benoted that in the MS community the currentlyaccepted unit for atomic or molecular mass is thedalton (Da), which is exactly one-twelfth themass of a 12C atom.

A mass spectrum is thus a plot of ion inten-sity as a function of mass/charge ratio (m/z). Toget the mass of an ion its charge must be known,but this can usually be determined from the spec-trum. Many molecules will fragment during theionization step, and their spectra then comprisea molecular ion and a series of fragment ions.Although such fragmentation will produce com-plicated spectra if many compounds are present,it nevertheless can provide valuable information

about the sub-structures present in the originalmolecules.

TOF mass analysers are velocity spectrom-eters, in that ions are separated by m/z basedupon different velocities acquired in an electro-static field. As all ions are accelerated throughthe same field, they all have the same nominalkinetic energy. Thus, their velocities will varyinversely with m/z. A mass spectrum is obtainedby monitoring ions arriving at the detector asa function of time. The defining relationshipthat equates drift time to m/z is given in

t vL

L V zm

21

/1 2

= = d dn n (3.1)

where t is the time for the ion to reach the detec-tor, L the length of the tube, v the ion velocityand V the accelerating voltage.

Early TOF analysers did not have particu-larly good resolution, generally less than about500, but their speed and virtually unlimited massrange made them popular. However, greatimprovements in the resolution of TOF spec-trometers followed from introduction of thereflectron (Fig. 3.6), which incorporates a seriesof electrical lenses that compensate for variationsin ion velocity (Cooper et al., 2005b). Modernreflectron TOF analysers are fast, which makesthem ideal for use as chromatographic detectors,and also have high mass resolving power andmass accuracy, a necessary requirement fordetecting molecules like inositol phosphates incomplex matrices.


Ion-storageregion

Field-free drift region Reflectron lenses

Detector

Deflectionplates

Fig. 3.6. Schematic representation of a time-of-flight (TOF) mass analyser with reflectrongeometry. Ions that enter the field-free drift regionmigrate to the detector at a rate that is dependenton their mass/charge ratio (m/z). The reflectronlenses compensate for variations in kineticenergies of the inject ions; these variations wouldotherwise produce broadened peaks and loss ofspectral resolution.

Electrospray Ionization MolecularMass Spectrometry of Inositol

Phosphates

Direct electrospray ionization time-of-flight mass spectrometry of inositol

phosphates

In previous studies of dissolved organic phos-phorus speciation in the Florida Evergladeswe noted that ESI was inefficient for organicphosphorus (Llewelyn et al., 2002). Figure 3.7includes ultrahigh-resolution ESI mass spectraof organic phosphorus standards before and

after the standards had been concentrated by aphosphorus-specific isolation procedure. Themass spectrum of the mixture before precipita-tion shows that many of these standards werenot seen as simple [M + H]+ ions. In addition,ESI efficiencies of organophosphates do notappear to be particularly high. These observa-tions further reinforce the need for selectiveconcentration of dissolved organic phosphorus.Fortunately, inspection of Fig. 3.7b suggests thatthe organic phosphorus isolation procedure weuse does not alter the ESI characteristics ofthese compounds.

We carried out a series of initial experi-ments using a ‘standard’ skimmer voltage setting


[AMP + H]+

[TP + H]+

[PEP + Na]+

[PEP + Na]+

[AMP + Na]+

[AMP + Na]+[AMP + H]+

250 300 350 400 450 500 550

[TP + H]+

[2RP + H]+

[2RP + H]+

250 300 350 400m/z

450 500 550(a)

(b)

Fig. 3.7. Positive ion ESI 9.4 T Fourier-transform ion-cyclotron resonance mass spectra showing thestandard dissolved organic phosphorus mixture – tyrosine phosphate (TP), adenosine monophosphate(AMP), phosphoenolpyruvate (PEP) and ribose phosphate (RP) – (a) prior to barium precipitation and(b) following barium precipitation. (From Llewelyn et al., 2002. Reprinted with permission from theAmerican Chemical Society.)

of 60 V. A JEOL AccuTOF mass spectrometer( JEOL, Inc., Tokyo, Japan) was used for all theMS measurements described here. This is a sin-gle-stage reflectron mass analyser with a mass res-olution of ~6000 and mass accuracy of less than5 ppm. This instrument includes an ESI sourcewith orthogonal geometry (Fig. 3.5b). Inositolphosphates were obtained from Sigma Chemicals(St Louis, Missouri, USA). Table 3.1 summarizesthe inositol phosphates used in these studies, aswell as molecular weights of the intact inositolphosphate and its molecular ion m/z value (M –H).

In the first experiment a mixture of sodiumsalts of myo-inositol monophosphate (C6H6(HnPO4)1; Mw = 260 Da) and myo-inositol hexak-isphosphate (C6H6(HnPO4)6; Mw = 660 Da) wasefficiently ionized, but we were surprised to

observe what appeared to be significant fragmen-tation of myo-inositol hexakisphosphate, with thebis- and pentakisphosphates also appearing in themass spectrum (Fig. 3.8). Although the presenceof lower-order inositol phosphates as impurities inthe myo-inositol hexakisphosphate standard can-not be ruled out, we believe that the phosphategroups in inositol phosphates may be relativelylabile during ESI, as the mass differences weobserve here (80 Da) represent the loss of HPO3group(s). This fragmentation process has beenobserved before with phosphate-containing pep-tides (Neubauer and Mann, 1999). Thus, intactmolecular ions may not always be the most abun-dant in the mass spectra of inositol phosphates.Nevertheless, these results are encouragingbecause they indicate that with sufficiently goodanalytical separation the molecular formulas of


Table 3.1. Isomeric forms of the six inositol phosphates used in this work, with molecular weights andm/z value at which the molecular ion of each appears in the electrospray ionization–time-of-flight(ESI–TOF) mass spectrum.

Molecular Molecular ion Inositol phosphate Isomer weight (Da) peak (Da)

IP1 myo-Inositol 2-monophosphate 260.0 259.0IP2 myo-Inositol 2,4-bisphosphate 340.0 339.0IP3 myo-Inositol 1,4,5-trisphosphate 420.0 419.0IP4 myo-Inositol 1,4,5,6-tetrakisphosphate 499.9 498.9IP5 myo-Inositol 1,3,4,5,6-pentakisphosphate 579.9 578.9IP6 myo-Inositol hexakisphosphate 659.9 658.9

m/z

400100 200 300 500 600 700 800

339 DaIP6;

659 Da498 Da

419 Da

259 Da

579 Da

Fig. 3.8. Electrospray ionization–time-of-flight (ESI–TOF) mass spectrum of myo-inositolhexakisphosphate (IP6); skimmer cone voltage 60 V.

the individual inositol phosphate species can beverified by ESI–MS. In addition, these resultssuggest that it may be possible to generate a com-mon ion from different inositol phosphates usingin-source fragmentation. This would allow phospho-rus-specific detection of peaks emerging fromHPLC separation columns.

These initial experiments were followed bya study to better understand the stability of inos-itol phosphates in the ESI source. This samemixture of myo-inositol monophosphate and myo-inositol hexakisphosphate was sprayed underidentical conditions, but varying skimmer volt-ages. The dramatic effect of skimmer voltage isevident in the mass spectra of Fig. 3.9. At 20 V(Fig. 3.9a) myo-inositol hexakisphosphate at 659Da is barely visible, while myo-inositol monophos-phate at 259 Da is quite intense. However,increasing the skimmer voltage to 80 V(Fig. 3.9b) greatly increases the myo-inositol hexa-kisphosphate molecular ion peak at 659 Da,while the myo-inositol monophosphate peak is notas intense as at 20 V. Figure 3.10 summarizesmolecular ion peak intensities over a wide rangeof skimmer voltages. Clearly, there is an opti-mum skimmer voltage between 40 and 80 V atwhich both molecular ion peaks are intense andsuitable for use in selected ion monitoring (SIM)to detect specific inositol phosphates. This con-clusion is confirmed by the mass spectrum of amixture of six inositol phosphates (Fig. 3.11) in

which the molecular ions (as M – H) of eachcompound are visible.

The low intensities of molecular ions at highskimmer voltages summarized in the data ofFig. 3.10 also suggest that it might be possible toidentify a fragmentation product common to allthe inositol phosphates and exploit in-sourcefragmentation for SIM after chromatographicseparation. One potential fragment appears inthe spectra of both myo-inositol monophosphateand myo-inositol hexakisphosphate at a skimmervoltage of 100 V at 79 Da and corresponds toPO3. This prominent fragment was previouslyobserved by Neubauer and Mann (1999) in astudy of phosphopeptide phosphorylation usingtriple quadrupole MS.

We therefore carried out another series ofexperiments in which we monitored the intensi-ties of molecular ions of all six inositol phos-phates and this 79 Da fragment at skimmer conevoltages of 20 and 60 V. Intensity ratios ofmolecular ion/fragment ion for the inositolphosphates are plotted in Fig. 3.12. From thesedata it is clear that all inositol phosphates frag-ment to a significant extent at 60 V. However,there is a very interesting trend at 20 V. The ten-dency to fragment, or at least to lose PO3,appears to increase as HPO3 groups are addedto the inositol backbone.

These results have somewhat negative conno-tations regarding the analysis of inositol phosphate


200

(a) (b)

400 600

m/z m/z

800 200 400 600 800

IP1;259 Da

IP6;659 Da

IP6;659 Da

IP1;259 Da

Fig. 3.9. ESI–TOF mass spectra of a myo-inositol monophosphate (IP1) and myo-inositolhexakisphosphate (IP6) mixture at skimmer cone voltages of (a) 20 V and (b) 80 V.

mixtures by direct ESI–MS. ESI of any inositolphosphate mixture will apparently produce molec-ular ions of each compound, but fragment ions willappear at the same m/z values as less phosphory-lated inositols. These results thus reinforce theneed for a moderately good analytical separationbefore the ESI–MS detection.

Size-exclusion chromatography withmass spectrometry detection of inositol

phosphates

Analysis of inositol phosphates by ESI–TOF–MSappears promising given the results describedabove. However, when attempting to detect inos-

itol phosphates in complex matrices such as soilsand animal manures it is necessary to isolatethem from other species that interfere with theirionization in the ESI source. This was one of theprimary findings of our work with organic phos-phates in an Everglades treatment wetland(Llewelyn et al., 2002): organic phosphorus doesnot compete very effectively for charge when ina high background of natural organic matter. Wethus began an evaluation of HP-SEC as a pre-liminary, on-line isolation step prior toESI–TOF–MS of inositol phosphates. SECwould appear to be the most useful liquid chro-matography separation technique for this pur-pose, as the mobile phases normally required areaqueous-based solutions with small amounts oforganic modifier(s) and volatile buffer salts.

Size-exclusion chromatography

SEC is based on a relatively simple principle:larger molecules are ‘excluded’ to a greaterextent from the inner spaces of a porous columnpacking material than smaller molecules, whichcan penetrate the openings of the small pores.The average residence time within the columnthus depends on the ‘effective’ molecular size, asmolecules in the pore volume of the packing areremoved form the flowing mobile-phase streamand do not move towards the end of the column.Retention in HP-SEC can be described by

V V VKr void pores= + (3.2)

where Vr is the retention volume of the solute(retention time × volumetric mobile-phase flow


200

5000

10,000

15,000

20,000

25,000

40

Cone voltage (V)

Inte

nsity

60 80 100 120

IP1

IP6

140

Fig. 3.10. Molecular ion peak intensities of myo-inositol monophosphate (IP1) and myo-inositolhexakisphosphate (IP6) as a function of skimmer cone voltage.

200 400

IP1;259 Da

IP3;419 Da

IP2;339 Da

m/z600 800

IP5;579 Da

IP6;659 Da

IP4;498 Da

Fig. 3.11. ESI–TOF mass spectrum of an inositolphosphate mixture at a skimmer cone voltageof 60 V.

rate), Vvoid is the volume of interparticle voidspace in the column, Vpores is the total pore vol-ume accessible to the smallest solutes and K is aconstant that describes the probability of thesolute penetrating into the pore space. The valueof K is a complex function of primarily molecularsize, but also factors such as shape and charge-density. Any molecule that is too large to enterany of the pores is totally excluded (K = 0), whilea small molecule can penetrate all the pores andhas access to the entire pore volume (K = 1). Size-exclusion columns are thus characterized byan exclusion limit, which is the molecular weightof all molecules with Vr = Vvoid , and total perme-ation limit, which is the molecular weight of allmolecules with Vr = Vvoid + Vpores. Molecules thatare of sizes that fall within exclusion and totalpermeation limits will be separated.

The separations described in this chapterwere carried out on a Beckman–Coulter SystemGold liquid chromatograph. Mobile phaseswere composed of 4:1 (v/v) mixtures ofwater/methanol. The water component of themobile phase was buffered with ammonium bicar-bonate at concentrations of 0.10 and 0.01 M. Weused a 250 × 4.6 cm i.d. PL-Aquagel-OH polymercolumn (Polymer Laboratories, Shropshire, UK),8 µm particle diameter, with a molecular weightseparation range of 100–30,000 Da. The columnwas calibrated with poly(styrene)sulphonate (PSS)

standards with nominal molecular weights of 1640,7900, 16,600 and 70,000 Da. Such standards areoften used to calibrate SEC columns for naturalorganic matter separations, as they are thought tobehave much like natural organic matter. Thechromatograms were characteristic of most HP-SEC separations: peaks are relatively broad andchromatographic resolution is not great, but mole-cules that are very different in size can be resolved.

SEC was performed using two mobile phasesthat differed only in their ionic strength. The firstconsisted of a mixture that was 80% water con-taining 0.10 M ammonium bicarbonate and 20%methanol. The second mobile phase was identical,except that the aqueous phase was only 0.01 M inammonium bicarbonate. Previous experiments byReemtsma and These (2003) demonstrated thatseparation of natural organic matter improved asthe ionic strength of the buffer increased, proba-bly due to ionic interactions between chargedsolutes and polar sites within the Aquagel polymermatrix. Calibration curves for the two mobilephases are included in Fig. 3.13. These curvesdemonstrate the classical log-linear relationshipbetween molecular weight and retention time inSEC. They also indicate that much better resolu-tion can be obtained with the higher ionicstrength mobile phase.

Our results confirm that high ionic strengthbuffers are necessary for efficient separation of


0IP1 IP2 IP3 IP4 IP5 IP6

10

20

Inte

nsity

rat

io

30

40

50

Molecular/fragment ionratio at 60 V

Molecular/fragment ionratio at 20 V

IP3 IP4 IP5 IP6

Inte

nsity

rat

io

4

00.5

11.5

22.5

33.5

Fig. 3.12. Ratios of intensities of inositol phosphate molecular ions to fragment ion peak at 79 Da andskimmer cone voltages of 20 and 60 V.

natural organic matter in size-exclusion polymercolumns. However, this presents a dilemma whenESI–MS will be used for detection, since highionic strength decreases signal intensity. Indeed,baselines in chromatograms using the 0.10 Mbuffer and ESI–TOF–MS detection were toonoisy to be useful. Thus, we were forced to tradethe chromatographic resolution obtainable athigh buffer strengths with acceptable signal inten-sities obtainable only at lower buffer strengths.

High-performance size-exclusionchromatography with ESI–TOF selected ion

monitoring mass spectrometry detectionof inositol phosphates

The behaviour of inositol phosphates in an ESIsource provides a number of opportunities fordetermining them in complex matrices. As notedpreviously, the SEC separation nicely isolates theinositol phosphates from much of the interferingnatural organic matter that would otherwiseobscure their ESI–MS signals due to ionizationsuppression. Although some compromise betweenseparation efficiency and signal intensity is neces-sary, acceptable isolation of inositol phosphatescan be achieved at lower mobile-phase ionicstrengths. Because inositol phosphates ionize atlower cone voltages primarily as molecular ions,individual inositol phosphates can be identified asthey elute from an SEC column by monitoring

the appropriate ion as a function of time. Thisapproach is generally referred to as SIM and isnow a well-established analytical technique. Ourapproach to SIM is to scan the entire mass rangeof interest repeatedly throughout a chromato-graphic separation. Then, a ‘mass chromatogram’can be reconstructed by plotting the intensity ofone molecular ion as a function of time.

SIM is demonstrated in Fig. 3.14. The totalion current chromatogram of a sample of myo-inos-itol bisphosphate shown in Fig. 3.14a was obtainedby summing the intensities of all ions within themass range being monitored, in this case 80–800Da. No peaks are visible because the IP2 signal isso low that it does not rise above the backgroundsignal produced by ionization of molecules in themobile phase. However, when the signal producedonly by the ion at m/z = 339 Da is plotted, themyo-inositol bisphosphate peak (as M – H) is clearlyvisible and intense relative to the background.Figure 3.14b is a classic example of a ‘mass chro-matogram’, in this case of inositol bisphosphate.

The previous experiments on directESI–TOF–MS indicated that all six inositol phos-phates we tested produced sufficiently intensemolecular ion peaks (as M – H) to be monitoredby this SIM technique. Figures 3.15 and 3.16include mass chromatograms for two of the sixinositol phosphates, separated as a mixture. Itshould be noted that both mass chromatogramswere obtained with only one size-exclusion sepa-


15

3

3.5

4 (b)(a)

4.5

log

Mw

5

17 19 21

Tr (min)

23 25

Fig. 3.13. Molecular weight (Mw) vs. retention time (Tr) calibration curves for poly(styrene)sulphonate(PSS) standards on a PL-Aquagel-OH polymer column with 8 µm particle diameter: (a) 80% watercontaining 0.10 M ammonium bicarbonate and 20% methanol; (b) 80% water containing 0.01 Mammonium bicarbonate and 20% methanol.

ration; the mass chromatograms were recon-structed from this single chromatogram by plot-ting the intensities of the appropriate masses. Alsoincluded in these figures are the mass spectra overthe entire mass range monitored (60–800 Da) atthe elution time of each compound. The rela-tively low background and lack of many non-inositol phosphate peaks in these spectra areno doubt one additional benefit of the HP-SECseparation prior to mass analysis.

High-performance size-exclusionchromatography with ESI–TOF massfragmentometry detection of inositol

phosphates

We previously identified a fragment at m/z 79that was common to all the inositol phosphates.This PO3 fragment could potentially be moni-tored in the SIM mode and serve as a signal forany inositol phosphate. This would convert theESI–TOF mass spectrometer into an inositolphosphate–specific detector, and accomplish whatwe previously achieved using ICP ionization and

elemental MS of phosphorus at m/z 31. This spe-cial approach to SIM in which a fragment ratherthan a molecular ion is monitored during a chro-matographic separation is sometimes referred toas ‘mass fragmentometry’, and the resulting chro-matogram, a ‘mass fragmentogram’.

A ‘mass fragmentogram’ of a mixture of allsix inositol phosphates is depicted in Fig. 3.17.This fragmentogram was reconstructed from con-tinued scanning of masses 60–800, and then plot-ting the intensity of the 79 Da peak as a functionof time. Surprisingly, this SEC column, with anominal separation range of 100–30,000 Da, wasable to resolve the inositol monophosphate (peakat 19 min) from the other inositol phosphates.However, myo-inositol bisphosphate through myo-inositol hexakisphosphate were not resolved.

Summary

ESI–TOF–MS offers several approaches to meas-uring inositol phosphates in complex environ-mental, agricultural and biological matrices. All


(a)

(b)5 10

Retention time (min)15 20

Fig. 3.14. High-performance size-exclusionchromatograms (HP-SEC) with ESI–TOFdetection of myo-inositol bisphosphate: (a) Totalion chromatogram; (b) selected ion monitoring(SIM) chromatogram at m/z 339.

5 10 15

Retention time (min)

20

100 200 300 400m/z

500 600 700 800

(a)

(b)

Fig. 3.15. HP-SEC and mass spectrum of myo-inositol monophosphate: (a) SIM chromatogram atm/z 259; (b) mass spectrum.

six inositol phosphates are sufficiently stable dur-ing the ionization process to be observed as (M – H) molecular ions. Unfortunately, they alsofragment to some extent, losing PO3 in a processyielding ions that appear at exactly the same m/zvalues as other less phosphorylated inositols.Thus, a physical separation by some sort of chro-

matography will be necessary before SIM of eachinositol phosphate can be quantitative.

However, the tendency to fragment can beused as an advantage, as all inositol phosphatesappear to produce PO3 as a common fragment ion.SIM of this fragment at 79 Da can thus convert theESI–TOF mass spectrometer into a phosphorus-specific chromatographic detector, in the same waythat inductively coupled plasma mass spectrometry(ICP–MS) monitoring of elemental phosphorus at31 Da has been used (Cooper et al., 2005a).

The combination of liquid chromatographyand MS would thus appear to offer a new, moresensitive and selective analytical method for quan-titatively identifying inositol phosphates in environ-mental samples. MS is significantly more sensitiverelative to NMR and also minimizes the effect ofother organic phosphates on the quantification ofindividual inositol phosphates. It has been notedthat many of the useful ion-exchange separationtechniques require pre-treatment of samples byhypobromite oxidation to remove other organicphosphates that co-elute and interfere with inositolphosphate quantification when phosphorus-specificdetectors that include post-column derivatizationare used (Irving and Cosgrove, 1981). These inter-fering phosphorus compounds would not be aproblem if MS with SIM were used as the chro-matographic detector, eliminating the need for apreliminary oxidation step. However, the ESIprocess that is critical to MS detection is highlysensitive to salts and polar compounds that com-pete with phosphates for charge in the ESI source.Thus, complex environmental samples will stillrequire extensive preliminary clean-up and isola-tion procedures to remove inositol phosphatesfrom metals and organics (e.g. high-molecularweight humic acids) to which they are boundbefore these sensitive and selective HPLC–ESI–MS techniques could be used.

Acknowledgements

This work was supported by grants from theSouth Florida Water Management District andthe United States Department of Agriculture(USDA). The assistance of Dr Umesh Goli,Director of the Mass Spectrometry Laboratoryat Florida State University Department ofChemistry, is greatly appreciated.


5(a)

10Retention time (min)

m/z(b)

100 200 300 400 500 600 700 800

15 20

10

Retention time (min)

15 20

Fig. 3.16. HP-SEC and mass spectrum of myo-inositol hexakisphosphate: (a) SIM chromatogramat m/z 659; (b) mass spectrum.

Fig. 3.17. HP-SEC SIM at m/z 79 (massfragmentogram) of an inositol phosphate mixture;skimmer cone voltage 60 V.

References

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Buscher, B.A.P., Irth, H., Andersson, E., Tjaden, U.R. and van der Greef, J. (1994) Determination of inositolphosphates in fermentation broth using capillary zone electrophoresis with indirect UV detection. Journal ofChromatography 678, 145–150.

Buscher, B.A.P., Vanderhoeven, R.A.M., Tjaden, U.R., Andersson, E. and Vandergreef, J. (1995) Analysis ofinositol phosphates and derivatives using capillary zone electrophoresis mass-spectrometry. Journal ofChromatography A 712, 235–243.

Casals, I., Villar, J.L. and Riera-Codina, M. (2002) A straightforward method for analysis of highly phosphory-lated inositols in blood cells by high-performance liquid chromatography. Analytical Biochemistry 300, 69–76.

Chen, Q.-C. and Li, B.W. (2003) Separation of phytic acid and other related inositol phosphates by high per-formance ion chromatography and its applications. Journal of Chromatography A 1018, 41–52.

Cooper, W.T., Llewelyn, J.M., Bennett, G.L. and Salters, V.J.M. (2005a) Mass spectrometry of natural organicphosphorus. Talanta 66, 348–358.

Cooper, W.T., Llewelyn, J.M., Bennett, G.L., Stenson, A.C. and Salters, V.J.M. (2005b) Organic phosphorus spe-ciation in natural waters by mass spectrometry. In: Turner, B.L., Frossard, E. and Baldwin, D.S. (eds) OrganicPhosphorus in the Environment. CAB International, Wallingford, UK, pp. 45–74.

Cosgrove, D.J. (1963) The chemical nature of soil organic phosphorus. I. Inositol phosphates. Australian Journal ofSoil Research 1, 203–214.

Cosgrove, D.J. (1966) Detection of isomers of phytic acid in some Scottish and California soils. Soil Science 102,42–43.

Cosgrove, D.J. (1969) Ion-exchange chromatography of inositol polyphosphates. Annals of the New York Academy ofSciences 165, 677–686.

Cosgrove, D.J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry and Physiology. Elsevier, Amsterdam, TheNetherlands.

Dragani, L.K., Berrie, C.P., Corda, D. and Rotilio, D. (2004) Analysis of glycerolphosphoinositol by liquid chro-matography–electrospray ionisation tandem mass spectrometry using β-cyclodextrin-bonded column. Journalof Chromatography B 802, 283–289.

Fenn, J.B., Mann, M., Meng, C.K., Wong, S.F. and Whitehouse, C.M. (1989) Electrospray ionization for mass-spectrometry of large biomolecules. Science 246, 64–71.

Harland, B.F., Smikle-Williams, S. and Oberleas, D. (2004) High performance liquid chromatography analysis ofphytate (IP6) in selected foods. Journal of Food Composition and Analysis 17, 227–233.

Henshall, A., Harrold, M.P. and Tso, J.M.Y. (1992) Separation of inositol phosphates by capillary electrophore-sis. Journal of Chromatography 608, 413–419.

Hsu, F.F. and Turk, J. (2000) Characterization of phosphotidylinositol, phosphatidylinositol-4-phosphate, andphosphotidylinositol-4,5-bisphosphate by electrospray ionization tandem mass spectrometry: a mechanisticstudy. Journal of the American Society for Mass Spectrometry 11, 986–999.

Hsu, F.F., Goldman, H.D. and Sherman, W.R. (1990) Thermospray liquid-chromatographic mass-spectrometricstudies with inositol phosphates. Biomedical and Environmental Mass Spectrometry 19, 597–600.

Irving, G.C. and Cosgrove, D.J. (1981) The use of hypobromite oxidation to evaluate two current methods forthe estimation of inositol polyphosphates in alkaline extracts of soils. Communications in Soil Science and PlantAnalysis 12, 495–509.

Irving, G.C. and Cosgrove, D.J. (1982) The use of gas–liquid chromatography to determine the proportions ofinositol isomers present as pentakis- and hexakisphosphates in alkaline extracts of soils. Communications in SoilScience and Plant Analysis 13, 957–967.

Kasim, A. and Edwards, H.M. (1998) The analysis for inositol phosphate forms in feed ingredients. Journal of theScience of Food and Agriculture 76, 1–9.

Kerwin, J.L., Tuininga, A.R. and Ericsson, L.H. (1994) Identification of molecular-species of glycerophospholipidsand sphingomyelin using electrospray ionization mass-spectrometry. Journal of Lipid Research 35, 1102–1114.

L’Annunziata, M.F. and Fuller, W.H. (1971) Nuclear magnetic resonance spectra of acetate derivatives of soil andplant inositol phosphates. Soil Science Society of America Proceedings 35, 655–658.

L’Annunziata, M.F. and Fuller, W.H. (1976) Evaluation of the mass spectral analysis of soil inositol, inositol phos-phates, and related compounds. Soil Science Society of America Journal 40, 672–678.


L’Annunziata, M.F., Gonzalez, J. and Olivares, L.A. (1977) Microbial epimerization of myo-inositol to chiro-inositolin soil. Soil Science Society of America Journal 41, 733–736.

Llewelyn, J.M., Landing, W.M., Marshall, A.G. and Cooper, W.T. (2002) Electrospray ionization Fourier trans-form ion cyclotron resonance mass spectrometry of dissolved organic phosphorus species in a treatment wet-land after selective isolation and concentration. Analytical Chemistry 74, 600–606.

McKercher, R.B. and Anderson, G. (1968a) Characterization of inositol penta- and hexaphosphate fractions of anumber of Canadian and Scottish soils. Journal of Soil Science 19, 302–310.

McKercher, R.B. and Anderson, G. (1968b) Content of inositol penta- and hexaphosphates in some Canadiansoils. Journal of Soil Science 19, 47–55.

Neubauer, G. and Mann, M. (1999) Mapping of phosphorylation sites of gel-isolated proteins by nanoelectrospraytandem mass spectrometry: potentials and limitations. Analytical Chemistry 71, 235–242.

Phillippy, B.Q. and Johnston, M.R. (1985) Determination of phytic acid in foods by ion chromatography withpost-column derivatization. Journal of Food Science 50, 541–542.

Reemtsma, T. and These, A. (2003) On-line coupling of size exclusion chromatography with electrospray ioniza-tion-tandem mass spectrometry for the analysis of aquatic fulvic and humic acids. Analytical Chemistry 75,1500–1507.

Sherman, W.R., Ackerman, K.E., Berger, R.A., Gish, B.G. and Zinbo, M. (1986) Analysis of inositol monophos-phates and polyphosphates by gas-chromatography mass-spectrometry and fast-atom-bombardment.Biomedical and Environmental Mass Spectrometry 13, 333–341.

Skoglund, E., Carlsson, N.-G. and Sandberg, A.-S. (1998) High-performance chromatographic separation of inos-itol phosphate isomers on strong anion exchange columns. Journal of Agricultural and Food Chemistry 46,1877–1882.

Steward, J.H. and Tate, M.E. (1971) Gel chromatography of soil organic phosphorus. Journal of Chromatography 60,75–78.

Taguchi, R., Hayakawa, J., Takeuchi, Y. and Ishida, M. (2000) Two-dimensional analysis of phospholipids bycapillary liquid chromatography/electrospray ionization mass spectrometry. Journal of Mass Spectrometry 35,953–966.

Turner, B.L., Papházy, M.J., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment.Philosophical Transactions of the Royal Society, London, Series B 357, 449–469.


4 Origins and BiochemicalTransformations of Inositol

Stereoisomers and Their Phosphorylated Derivatives

in Soil

Michael F. L’AnnunziataThe Montague Group, PO Box 5033, Oceanside,

CA 92052-5033, USA


The determination of inositol phosphatesin environmental samples is an analytical challenge that has hampered research on thistopic for decades. The role of inositol phos-phates in the environment, including the originsand biochemical transformations of the soilinositol phosphates and their detection by mod-ern instrumental and isotopic techniques, wasreviewed recently by Turner et al. (2002), anddetailed reviews of the use of nuclear mag-netic resonance (NMR) spectroscopy and massspectrometry for the analysis of inositol phos-phates in environmental samples also appear inthis volume (see Murthy, Chapter 2, andCooper et al., Chapter 3). This chapter summa-rizes early work (1967–1969) on the extractionand characterization of inositol phosphates insoil using a range of analytical procedures,including electron-impact mass spectrometry,infrared spectroscopy and proton NMR spec-troscopy. It also describes subsequent work(1974–1977) conducted using isotopic-labellingtechniques, and concludes with ideas on newapproaches to research on the inositol stereoiso-mers and their phosphorylated derivatives insoils and other environmental samples using sta-ble and radioactive isotopes.

Development of Methodologies forCharacterization of Inositol

Phosphates in Soil

Extraction and preparativechromatography

Inositol phosphates were extracted from leaf litteron the soil surface and the underlying soilorganic matter of a forest and desert soil by themethod of Anderson (1956). A flow chart of theextraction method is illustrated in Fig. 4.1. Theprecipitated fractions of extractable phosphorus-labelled precipitates A, B and C of Fig. 4.1 werecombined, the phosphate groups were removedby hydrolysis in hydrochloric acid, the solutiondeionized with exchange resin and finally con-centrated by evaporation to 25 ml.

The phosphate groups were removed fromthe inositol rings to facilitate the separation andisolation of the inositol stereoisomers by prepara-tive paper chromatography and their subsequentrecrystallization and structural identification by arange of analytical techniques (L’Annunziata,1970; L’Annunziata and Fuller, 1971a;L’Annunziata et al., 1972). The objective of the

study was to investigate the origins of the soilinositol phosphate stereoisomers by assessing thestereochemistry of inositols in leaf litter on thesoil surface and in the soil organic matter of twosoil types, a desert and a forest. The extractedphosphates were subjected to acid hydrolysis toremove all phosphate groups: a major portion ofthe hydrochloric acid was removed by vacuumdistillation, anions and cations were removed bywashing the extract through a column of deioniz-ing resin and the aqueous solution concentratedto 25 ml by evaporation in an open beaker on ahot plate at moderate heat (60–80ºC). The inosi-tols in the solution were separated by descendingpreparative paper chromatography by spreading~0.4 ml of the solution onto individual large

(56 × 23 cm) chromatogram sheets under thewarm air of a hairdryer. Approximately 50 chro-matograms were required to isolate all of theinositol stereoisomers in the 25 ml concentratedextract. More details on the extraction and chro-matographic procedures are available inL’Annunziata (1970) and L’Annunziata andFuller (1971a). The inositol stereoisomers wereeluted from the paper with hot deionized waterand recrystallized in aqueous acetone or ethanolwhen possible (i.e. whenever sufficient samplewas obtained). The inositol stereoisomers wereidentified by electron-impact mass spectrometry,infrared spectroscopy and solution 1H-NMRspectroscopy of the free inositols and their hexac-etate derivatives.

42 M.F. L’Annunziata

Soil

Soil

Supernatant

Supernatant

Precipitate A75% of extractable P

Precipitate B (Ba phosphates)

Precipitate C (Ba phosphates)

Vacuum distillation to concentrate

Supernatant

2 l of 10% BaAc and centrifuge

Supernatant

Supernatant

Supernatant

1. Precipitates A, B, C combined2. Refluxed 2 days with 1 l of 1:1 conc. HCI3. Deionized with exchange resin4. Evaporated to 25 ml volume

Wash with 0.2 M HCIExtract with 1 M NaOHCentrifuge

Wash with 1 M NaOHCentrifuge

200 ml HAcAcidify with HCI to pH 0.5Centrifuge

Humic acid (discard)5% of extractable P

Raise pH to 8.5 with NH4OHCentrifuge

Fig. 4.1. Flow chart illustrating the method of Anderson (1956) used to extract phosphorus from soil andplant samples. When used to extract phosphorus and its compounds from leaf litter, the word ‘soil’ in theflow chart should be replaced with ‘plant matter’.

Electron-impact mass spectrometry

The electron-impact mass spectra of the inositolstereoisomers (myo- and D-chiro-inositol) wereidentical as expected, but the mass spectra pro-vided initial evidence for the basic inositol struc-ture by the molecular ion peak at m/e 180, thecharacteristic base peak at m/e 73 (C3H5O2

+) andother characteristic ion peaks at m/e 102(C4H6O3

+) and m/e 60 (C2H4O2+). These are dis-

cussed in more detail below. The electron-impactmass spectrometers available at the time causedstrong ionization and fragmentation and conse-quently a weak molecular ion peak. However,the fragmentation pattern together with themolecular ion peak was indispensable in makingunequivocal identification of the compound asinositol or one of its derivatives. The mass spec-tral fragmentation of inositol, inosose and inosi-tol-related compounds were studied in detail, themolecular formulas of the major ion peaks deter-mined and the mass spectral ion fragmentationscheme postulated (L’Annunziata, 1970;L’Annunziata and Fuller, 1976). The mass differ-ence of the major ion fragments was demon-strated to be a useful indicator of specificstructural changes of the inositols.

Typical electron-impact mass spectraobtained for the inositol stereoisomers (myo- andD-chiro-) and their natural products DL-epi-inososeand quebrachitol (monomethyl ether of L-chiro-inositol) are illustrated in Fig. 4.2. A detailed studywas made to identify the mass spectral ion frag-ments and the fragmentation scheme of the inosi-tols by means of mass spectral peak shiftsthat resulted following deuterium labelling ofthe hydroxyl groups of the inositol, inososeand monomethyl ether (L’Annunziata, 1970;L’Annunziata and Fuller, 1976). The structures ofthe mass spectral ion fragments were postulatedusing the molecular formulas of the fragment ionpeaks in conjunction with evidence from peakshifts produced by the mass spectra of molecularderivatives of inositol (e.g. inosose, quebrachitoland inositol hexacetate) and the deuterium-labelled analogues of the derivatives. The molecu-lar formulas were obtained by high-resolutionmass spectrometry, or postulated for ion frag-ments of low mass by the elimination of unlikelycombinations of carbon, hydrogen and oxygenlisted in mass and abundance tables (Beynon andWilliams, 1963; Buchs et al., 1968; L’Annunziata,

1970). The major mass spectra fragment ionsand fragmentation pathway of myo-inositol is illustrated in Fig. 4.3 (L’Annunziata, 1970;L’Annunziata and Fuller, 1976).

The mass spectra of the DL-epi-inososehelped to elucidate the mass spectral fragmenta-tion scheme of myo-inositol, but its mass spectraalso serve as a useful future reference for theidentification of the inosose in soil systems,because the cyclic ketone stereoisomers havebeen previously proposed to occur as phosphory-lated and non-phosphorylated intermediates insoil inositol and inositol phosphate epimerizationreactions (e.g. L’Annunziata and Fuller, 1971a;Cosgrove, 1972; L’Annunziata, 1975).

The ion fragment of m/e 163 observed inthe mass spectrum of myo-inositol was of lowintensity (0.1% of base peak); however, the differ-ence in mass between this ion fragment and theprotonated molecular ion of m/e 181 (Fig. 4.2a)was 18 mass units. This is equivalent to the lossof a water molecule, a common initial fragmenta-tion pathway for polyhydroxy compounds. Theion fragment at m/e 144 is equivalent to the lossof 36 mass units or two water molecules. Themechanism for this elimination of water remainsuncertain (Reed et al., 1962), although the threepossible 1,2-, 1,3- and 1,4-eliminations may beconsidered. Two consecutive 1,2,-eliminationsare illustrated in Fig. 4.3, resulting in the forma-tion of ions II and III of m/e 163 and 144,respectively.

A peak shift of two mass units from m/e 144(C6H8O4

+) to m/e 146 occurred in the mass spec-trum of myo-inositol after deuterium labelling ofthe hydroxyl groups (C6H6(OD)6 of Fig. 4.2b),indicating the presence of two hydroxyl protonson the ion of m/e 144. This was explained by aketo–enol conversion between ions III and IV(Fig. 4.3) in which ion IV would shift to m/e 146for the deuterium-labelled inositol. A major ionfragment at m/e 102 (C4H6O3

+) contains two orthree hydroxyl groups as indicated by its shift tom/e 104 and 105 after deuterium labelling(Fig. 4.2b). The formation of the fragment ion atm/e 102 (ion V of Fig. 4.3) may be expressed bythe loss of a neutral ketene molecule from the ionof m/e 144 (ion IV of Fig. 4.3). The loss of keteneis characteristic of cyclic ketones. The keto–enolforms of ions V and VI of m/e 102 (Fig. 4.3) canexplain peak shifts of two and three mass units ofthe deuterium-labelled hydroxyl groups.

Inositol Stereoisomers 43


Fig. 4.2. Electron-impact mass spectra of (a) myo-inositol; (b) myo-inositol-d6, C6H6(OD)6; (c) DL-epi-inosose; and (d) quebrachitol. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1976.)

HH

H

100

90

80

70

60

50

% o

f bas

e pe

ak%

of b

ase

peak

% o

f bas

e pe

ak%

of b

ase

peak

40

30

20

10

100

90

80

70

60

50

40

30

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10

100

90

80

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20

10

20 30 40 50 60

60 (C2H4O2+)

62 (C2H2D2O2+)

60 (C2H4O2+)

73 (C3H5O2+) 87 (C4H7O2

+)

102 (C4H6O3+)

116 (C5H8O3+)

144(C6H6O4+)

85 (C4H5O2+)

104 (C4H4D2O3+) 105 (C4H3D3O3

+) 146 (C6H6D2O4+)

75 (C3H3D2O2+)

73 (C3H5O2+)

102 (C4H6O3+)

70 80 90 100 110

myo -Inositol,C6H6(OH)6

myo -Inositol-d6,C6H6 (OD)6

120(a)

(b)

(c)

(d)

m/e 163: 0.1%m/e 181: 0.1%

m/e 179: 1.5% (MH+)m/e 160: 2.4% (M -+H2O)

130 140

20 30 40

OH

OH

CH3OOH

OHOH

H

H

HH

H

H

50 60 70 80 90 100 110

Quebrachitol,

m/e 158: 3.5 %m/e 194: 0.1%m/e 195: 0.2%

C6H6(OH)5 OCH3

120 130 140

20 30 40 50 60 70 80

m/e

90 100 110 120 130 140 150

142 (M−+2 H2O)

20 30 40 50 60

60

73

71

HC C C

10289

H

D,L-epi -Inosose,C6H5O (OH)5

+

OH

OH

HO

70 80 90 100 110 120 130 140 150

144 (C6H8O4+)

H

H

H

OHOH

HH

H

H

+

=O

HOH

HO

HOOH

OH

OH

OHOH

OH

O=C=C−C +H H

OH

O=C−C=CH H

OH

. .

. .

..

The base peak (most intense peak) at m/e73 (C3H5O2

+, Fig. 4.2a) in the mass spectra ofinositol is produced by an ion fragment with twohydroxyl groups indicated by its shift to m/e 75after deuterium labelling as illustrated in

Fig. 4.2b. The ion is illustrated as the ring struc-ture VII of Fig. 4.3 or as the resonance-stabilizedallylic ion VII' in Fig. 4.4. This ion fragment(VII or VII') is also responsible for the base peakin the mass spectra of inosose and quebrachitol.


OH

I, m/e 181 II, m/e 163 III, m/e 144

IV, m/e 144

VIII, m/e 60VII, m/e 73

V, m/e 102VI, m/e 102

H

−CO

−H

+

H H

H H

C C−CH•

OH OHHOHO

OH OH

OH

OH

O

O

−CH2 = C = O

OH

−CH2 = C = O

O

OH

HO

OH

−H2OOH

OH

+

+ + +

+

HO

HO

HO

OH

OH

OH

OH

OH

OH

OH

+ +

−H2O

OH

Fig. 4.3. The major electron-impact mass spectral fragmentation ions and fragmentation pathway ofmyo-inositol. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1976.)

H

C CH

H

OH

H

H O C C C+

OCH3

H

X′, m/e 85

O+ C CH H O

OH OH

HO

XI, m/e 116 XII, m/e 116

OCH3 OCH3C

OCH3

H

O

X, m/e 85

OCH3

+

VII′, m/e 73

OHHO

IX, m/e 87 IX′, m/e 87

H

H+

H

H

C CH

C

OCH3

H+

+

HOOCH3

HO

HHH+

CCCC+

HO

(a) (b)

(d)(c)

. ... . .

Fig. 4.4. Major ion fragments (a) base peak at m/e 73 (C3H5O2+) of the electron-impact mass

spectrometry of myo-inositol, DL-epi-inosose and quebrachitol; (b) the second most intense peak at m/e87 (C4H7O2

+) at 98.9% of the base peak from the mass spectrum of quebrachitol; (c) the third mostintense peak at m/e 85 (C4H5O2

+) at 83% of the base peak from the mass spectrum of quebrachitol; and(d) an intense peak at m/e 116 (C5H8O3

+) from the mass spectrum of quebrachitol. (From L’Annunziata,1970; L’Annunziata and Fuller, 1976.)

The second-most predominate peak in the massspectra of inositol and inosose and also a pre-dominant peak in the mass spectra of quebrachi-tol is that at m/e 60 (C2H4O2

+), which containstwo hydroxyl groups (ion VIII of Fig. 4.3) asevidenced by its shift to m/e 62 followingdeuterium labelling of the hydroxyl groups.The mass spectra of quebrachitol labelledwith deuterium in the hydroxyl groups[C6H6(OD5)OCH3] and of the hexacetate deriv-atives of myo-inositol [C6H6(OAc)6] and quebra-chitol [C6H6(OAc5)OCH3] provided furtherevidence for the ion fragments and fragmenta-tion scheme described (L’Annunziata, 1970;L’Annunziata and Fuller, 1976).

Infrared spectroscopy

The infrared spectra provided perfect finger-prints for the myo- and chiro-inositol stereoisomersat every absorption frequency (e.g. for the pon-derosa pine needles and forest soils; Fig. 4.5).The infrared spectra can serve as a tool in distin-guishing specific inositol stereoisomers as evi-denced by the dissimilarities between spectra ofthe myo- and chiro-inositol standards. These occurprimarily in the ‘fingerprint’ region between1350 and 800 cm�1. For example, marked dis-similarities are evident at 900 and 930 cm�1 formyo-inositol (left panel of Fig. 4.5) and at 870 and905 cm�1 for chiro-inositol (right panel of Fig. 4.5).These may be due to differences in C–H rockingfrequencies (Dyer, 1965). Also, marked dissimi-larities are seen by four relatively sharp absorp-tion frequencies at 1000, 1075, 1110 and 1150cm�1 in the infrared spectrum of myo-inositol,which appear as two broad absorption frequen-cies at 1010 and 1100 cm�1 in the infrared spec-trum of chiro-inositol. These are assigned todifferences in C–O stretching frequencies(Lambert et al., 1998). The strong absorptionsassigned to O–H stretching frequencies at 3300cm�1 and the weaker absorption at 2900 cm�1

assigned to C–H stretching frequencies werecommon in all inositol infrared spectra studied.The very sharp truncated absorption peak at1600 per cm seen in all the infrared spectra isthat of a 0.07 mm polystyrene film used as a ref-erence standard.

Proton nuclear magnetic resonancespectroscopy

The 1H-NMR spectra provided strong evidencefor the existence of inositol stereoisomers(Fig. 4.6). The most powerful spectrometers avail-able to us at the time were 60 and 100 MHzinstruments, so relatively large samples (>5 mg)were needed to obtain spectra in deuterium oxidewith specially designed micro-NMR tubes. Themyo-inositol standard and myo-inositols isolatedfrom plant litter or underlying soil gave NMRspectra similar to those illustrated in the leftpanel of Fig. 4.6, with complex chemical shifts at5.99, 6.31, 6.40, 6.49, 6.59, 6.70 and 6.78τ.Spectra of the chiro-inositol standard and thechiro-inositols isolated from plant litter and under-lying soil differed from the myo-inositol spectrumby yielding complex chemical shifts at 5.97, 6.00,6.31, 6.33, 6.38 and 6.42τ at different relativeintensities than those for myo-inositol (right panelof Fig. 4.6).

The 1H-NMR spectra of the hexacetatederivatives of the myo- and chiro-inositols isolatedfrom the soil and plant litter were also perfect fin-gerprints of authentic myo- and chiro-inositol(L’Annunziata, 1970; L’Annunziata and Fuller,1971b). Up to 88% yields were obtained with theacetylation of inositol samples as small as 20 mg.The increase in the molecular weight of the inos-itol upon acetylation was advantageous due tothe very small amounts of inositols isolated fromplant or soil. The NMR spectra of the acetatederivatives (not illustrated here) permitted theobservation of the number of axial and equatorialacetoxy groups as sharp peaks due to the acetoxyprotons (CH3CO2–) lacking the complexities pro-duced by proton spin coupling.

Identification of D-chiro-inositol

The optical rotation of the chiro-inositol enan-tiomer from the desert soil–plant system wasdetermined to be +56º measured by a recordingpolarimeter (L’Annunziata, 1970). At about thesame time, Cosgrove (1969) also reported the D-chiro-inositol as a hexakisphosphate. Further evi-dence of the D-chiro-inositol was subsequentlyprovided by measurement of the positive optical


Inositol Stereoisom

ers47

30.0

0.10

0.20

0.300.400.500.600.701.0

4000 3000 2000 1500 1200 1000

cm−1

900 800

Inositol isomerfrom the forest soil

Inositol isomerfrom P. ponderosa needles

700

4000 3000 2000 1500 1200 1000 900 800 700

4000

INFRACORD � 1371282

INFRACORD � 1371282

3000 2000 1500 1200 1000 900 800 700

1.5

4 5A

bsor

banc

e6 7

Wavelength (microns)

8 9 10 11 12 13 14 15 3 4 5 6 7



8 9 10 11 12 13 14 15

3 4 5 6 7 8 9 10 11 12 13 14 15

∞

0.00.0

0.10

0.20

Abs

orba

nce

Abs

orba

nce

0.30

0.40

0.500.600.70

1.01.5

0.10

0.20

0.300.400.500.600.701.01.5∞

∞

0.0

0.10

0.20

0.30

0.400.500.600.70

1.01.5∞

0.0

0.10

0.20

0.30

0.400.500.600.70

1.01.5∞

0.0

0.10

0.20

0.30

0.40

0.500.600.70

1.01.5∞

(b)

(c)

(a) L-chiro-Inositol

Inositol isomerfrom chromatographic spot b2

(a)

cm−1

cm−1

myo-Inositol

(b)

Fig. 4.5. Characteristic infrared spectra of myo- and chiro-inositol. The left panel shows spectra of myo-inositol obtained from (a) a standard sample and (b) purified samples from a forest soil and (c) ponderosa pine needles from the soil surface. The right panel shows spectra of (a) a chiro-inositol standard and(b) the chiro-inositol isomer isolated from a desert soil or its velvet mesquite leaf litter on the soil surface. Regardless of source (i.e. plant or soil) the infraredspectra of pure myo-inositol or chiro-inositol were exact fingerprints. (From L’Annunziata, 1970; L’Annunziata and Fuller, 1971a; L’Annunziata et al., 1972.)

48M

.F.L’Annunziata

100200 6.49τ

5.99t6.40

6.70t6.78t

DDS

6.31t6.59t

H2O

300

(a) (a)

400 100200300400

8.0 7.0 6.0 5.0 4.0 3.0

6.42t

6.31t 6.33t

6.38t

6.00t5.97t

H2O

2.0 1.0 0

2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10

100050025010050

PPM(t)

H

HH H

H

HOH

OH

OH

OH

OH

HO

L-chiro-inositol

myo - InositolH

HO

HO

HO

H

H H

H

H

OH

OH

OHD

HHz D

HCPS

PPM(d)

8.0

DDS

100200300400

(b) (b)

100200300400100050025010050

1000

2.0 3.0 4.0 5.0 6.0 7.0 8.0 9.0 10

50025010050

6.49t

Forest soil inositol

6.70τ5.99τ

H2O

7.0 6.0 5.0 4.0PPM(δ) 3.0

TMS

H2O

2.0

Inositol isomerfrom chromatographic spot b2

1.0 0

DH

HzD

HCPS

PPM(t)

100050025010050

Fig. 4.6. Characteristic proton nuclear magnetic resonance (NMR) spectra of myo- and chiro-inositol. The left panel shows 100 MHz NMR spectra of myo-inositol obtained from (a) a standard sample and (b) a purified sample from a forest soil. Peaks due to DDS internal standard are marked. The right panelshows 60 MHz NMR spectra of (a) a chiro-inositol standard and (b) a chiro-inositol isomer isolated from the soil or its leaf litter on the soil surface. Regardlessof source (i.e. plant or soil) the NMR spectra of pure myo-inositol or chiro-inositol were exact fingerprints. (From L’Annunziata, 1970; L’Annunziata and Fuller,1971a.)

rotation of the chiro-inositol isolated from the for-est soil (L’Annunziata et al., 1972).

Identification of muco-inositol

Another inositol stereoisomer, identified as muco-inositol, was isolated from velvet mesquite leaf lit-ter on the soil surface, but was not identified inthe desert soil. The basic inositol structure wasconfirmed by the characteristic infrared spectraof the inositol and of its hexacetate derivative. Nostandard NMR spectrum was available for theunequivocal identification of muco-inositol, but itsmajor 1H-NMR absorption peak at 1.53 ppmwas in very close agreement to the 1.52 ppm formuco-inositol reported by Brownstein (1959). Themelting point (m.p.) of the hexacetate derivative(175.5–177ºC) was in close agreement with thatof authentic muco-inositol hexacetate (177–178ºC)reported by Nakajima et al. (1959). muco-Inositoloccurs in higher plants (Richter et al., 1990;Wanek and Richter, 1995; Peterbauer andRichter, 1998), but neither muco-inositol nor itsphosphorylated form has been reported in soil.Further discussion of the inositol phosphatestereoisomers can be found elsewhere (Turner,Chapter 12, this volume).

Significance of InositolStereoisomers in Soil

Not all of the inositol stereoisomers present in thesoil or plant samples were identified, due to thesmall amounts isolated and the limited power ofthe NMR spectrometers during the 1960s. scyllo-Inositol and neo-inositol phosphates occur in soils(Turner et al., 2002) and may have been presentin our samples, as indicated by a small amount ofan unidentified compound in one of the paperchromatogram bands (see c and d of Fig. 6 inL’Annunziata, 1970, p. 38). However, it was notpossible to recrystallize and run NMR spectra onthat small sample.

All chromatographic separations and massspectrometry and NMR spectroscopic measure-ments were performed on acid hydrolysates ofthe phosphate fractions A, B and C of plant orsoil extracts according to the method ofAnderson (1956) illustrated in Fig. 4.1. Hydrolysis

of the phosphates was performed by refluxing theprecipitates A, B and C in 1:1 hydrochloric acidfor 2 days, followed by deionization withexchange resin to remove inorganic ions to yieldaqueous solutions of the free inositols. At the timeof these studies (1967–1969) only electron-impactmass spectrometry was available, which requiredsamples in crystalline form that could bevolatilized with heating in the mass spectrometerfollowed by electron impact with molecules in thevapour phase. Consequently, mass spectrometryof inositols in the phosphorylated form was notpossible, so the phosphate groups were removedto facilitate identification of the inositol stereoiso-mers. Likewise, only 60 or 100 MHz NMR spec-trometers were available, which required thedissolution of the pure crystalline inositol in deu-terium oxide. The stability of the inositols underacid hydrolysis was tested by analysing myo-inosi-tol by 1H-NMR spectroscopy before and aftertreatment with 20% deuterated hydrochloric acidunder reflux, but no observable change occurredafter 3 days.

The studies described above were intendedto investigate only the organic carbon stereo-chemistry (inositol moiety) of the phosphates bymass spectrometry and NMR spectrometry.Subsequently, Cosgrove (1980) reported that con-siderable amounts of polysaccharide and nitroge-nous material could exist in the precipitatefractions isolated by the method of Anderson(1956), raising the possibility that some unphos-phorylated inositols may be extracted along withthe phosphorylated forms, particularly in extractsof plant matter. Ion-exchange chromatographicseparation of inositol phosphate fractions prior toacid hydrolysis would therefore be required toconclude irrefutably that the inositols are isolatedonly in the phosphorylated form.

The fact that both the myo-inositol andD-chiro-inositol were identified in the plant litteron the soil surface and in the underlying soilpointed to plant residue as a possible origin of thesoil inositol phosphate stereoisomers. However,soil microbial epimerization of myo-inositol toD-chiro-inositol was suggested as another possibleorigin of this stereoisomer (L’Annunziata, 1970)in light of the only structural difference betweenthe two stereoisomers, which is the stereochem-istry at a single carbon atom (L’Annunziata,1970; Fig. 4.7). The epimerization of myo-inositolto D-chiro-inositol involving a cyclic ketone


intermediate in Trifolium incarnatum was observedby Scholda et al. (1964), but there was no evi-dence then of any such reaction occurring insoils. Subsequent work was therefore undertakenwith the radioisotope 14C to provide unequivocalevidence for the soil microbial epimerization ofmyo-inositol to D-chiro-inositol.

Isotopic Techniques for StudyingInositol Phosphates in Soil

When myo-inositol uniformly labelled with 14C[14C(U)-myo-inositol] became available,L’Annunziata et al. (1975) proposed that it couldbe used to elucidate the pathways of myo-inositoltransformations in soil. The only stereoisomers ofinositol known to occur in soils as phosphory-lated forms were the myo-, D-chiro-, neo- and scyllo-inositol. The latter three differ from myo-inositol(the most abundant of the isomers in soil, plantand animal systems in phosphorylated or freeform) only at one carbon atom (Fig. 4.8). Thissuggested the possibility that D-chiro-, neo- andscyllo-inositol or their phosphates originated insoils by the epimerization of myo-inositol or itsphosphate at one of the carbon atoms marked bysymbols in Fig. 4.8.

To find the fate of myo-inositol and its hexak-isphosphate in soils, studies were initiated with 14Cisotope tracer. A study of the soil microbial metab-olism of uniformly labelled 14C(U)-myo-inositol,14C(U)-myo-inositol hexakisphosphate and 14C(U)-iron(III) phytate was carried out. The labelledcompounds were incubated in either microbiallyactive or sterilized forest soil in Bartha and Pramer(1965) incubation flasks (Fig. 4.9). The soil wastaken from the A1 horizon of a red forest soil

(Andisol), had a pH of 5.8 and contained 10.9%organic matter and 18% clay (L’Annunziata andGonzalez, 1977). The flasks permitted a determi-nation of the soil microbial metabolism rates bymeasurement of the evolution rates of 14CO2trapped by 0.1 M KOH in an attached side arm.The results indicated that 61% of the carbon inmyo-inositol, 1.9% of the carbon in myo-inositolhexakisphosphate and none of the carbon iniron(III) phytate were oxidized to carbon dioxideon 12 days of incubation (Table 4.1; L’Annunziataand Gonzalez, 1977). No carbon was oxidized to


1 2

3

45

myo -Inositol(1,2,3,5/4,6)-Inositol

neo -Inositol(1,2,3/4,5,6)-Inositol

scyllo-Inositol(1,3,5/2,4,6)-Inositol

D-chiro-Inositol(1,2,4/3,5,6)-Inositol

6

12

3

45

6

12

3

45

6

1 2

3

45

6

Fig. 4.8. The soil inositol stereoisomers. Thestereochemical differences of the D-chiro-, neo- andscyllo-inositol stereoisomers or their phosphates incomparison with the myo-inositol stereoisomer areindicated by the symbols ●, ■ and ▲, respectively.(Adapted from L’Annunziata, 1975.)

1 2

3 3 O 3

45

6

2 1H

−

4

Inosose intermediate

5

21

6

4

myo-Inositol(1,2,3,5/4,6)-Inositol (D)-chiro-Inositol

(1,2,4/3,5,6)-Inositol

5

6

Fig. 4.7. The proposed microbial epimerization of myo-inositol to D-chiro-inositol in soil. (FromL’Annunziata, 1970.) Numbering follows International Union of Pure and Applied Chemistry (IUPAC)nomenclature.

The rate of oxidation of 14C-labelled myo-inositolhexakisphosphate to carbon dioxide droppedrapidly from 0.99% after day 1 to 0.17% after day2, and then to 0.05% after day 12 (Table 4.1),indicating a rapid fixation of this compound

into a form unavailable to microorganisms(L’Annunziata and Gonzalez, 1977).

The soil incubated with 14C-myo-inositolunder normal and sterile conditions was fraction-ated according to Fig. 4.1. The quantities of 14Cfound in each soil fraction for both the non-ster-ile and sterile soils are shown in Table 4.2(L’Annunziata and Gonzalez, 1977), which liststhe percentage of the total radiocarbon remain-ing in the soil after 60% and 0% of the radioiso-tope label had evolved as 14CO2 for thenon-sterile and sterile treatments, respectively.The quantity of 14C encountered in each fractionis greatly dependent upon non-sterile or sterilesoil conditions.

As indicated in Table 4.2, fractionation of thesoil incubated with 14C(U)-myo-inositol under non-sterile conditions yielded small quantities ofradioisotope in the water- and HCl-extractablefractions. Larger quantities of 14C were encoun-tered in the unextractable and the humic-acid frac-tion. The unextractable fraction may consistof clay-bound inositol and its metabolites. Thephosphate fractions E, F and G were previouslyshown to contain myo-inositol hexakispho-sphate (Anderson, 1956; L’Annunziata, 1970;L’Annunziata and Fuller, 1971a). Phosphate frac-tions E, F and G of Table 4.2 (notation used in thewriter’s paper of 1977) are the same as fractions A,B and C of Fig. 4.1. Significant quantities of 14Cwere found in each of these fractions. Summing thepercentages of the 14C in each of the fractions withthe value of 61.0% for the radiolabel evolved as14CO2 (Table 4.1) yielded a total of 100.72%. Thisaccounts quantitatively for all of the 14C(U)-myo-inositol applied to the non-sterile soil. The phos-phate fractions E, F and G were submitted tohydrochloric acid hydrolysis to remove the phos-phate groups, and 14C-myo-inositol was identified inthe hydrolysed-phosphate fractions.

The soil-bound inositol carbon predomi-nates under non-sterile conditions (Table 4.2).The adsorption of inositol by clay minerals hasbeen demonstrated (Greenland, 1956). The highmineral adsorption of 14C (31.5% soil-bound orunextractable) in the non-sterile soil is consid-ered to be caused by the adsorption of microbialmetabolites. The predominance of metabolites of14C-myo-inositol in the non-sterile soil was esti-mated on the basis of the relatively high evolu-tion of 14CO2 under non-sterile conditions. Thehumic acid-bound inositol carbon formed under


i

c

g

h

f

b

a

d

e

Fig. 4.9. A Bartha and Pramer incubation flask.The components and contents of the flask are asfollows: (a) 50 g soil moistened to 70% fieldcapacity or other suitable moisture content; (b) 250ml Erlenmeyer flask fused to (c) a 50 ml test tubewith round bottom; (d) Ascarite (NaOH-coatedsilica) absorber of atmospheric carbon dioxide;(e) rubber stopper; (f) stopcock to permit incomingair when opened and (e) rubber stopper removedduring sampling of KOH; (g) 15 cm long 15-Gneedle with short length of polyethylene tubingextending to the bottom of the round base of theside arm; (h) rubber policeman cap, which isreplaced with a calibrated syringe to allow for theinjection of (i) 10 ml of 0.1 M KOH solution into theside arm and to allow for the removal of the KOHsolution and exchange with fresh KOH solution forthe measurement of 14CO2 at periodic intervals.(From Bartha and Pramer, 1965. Reprinted withpermission from Lippincott Williams & Wilkins.)

non-sterile soil conditions was abundant,although only a trace was encountered in thesterile soil (Table 4.2). The low level of 14C inthe humic acid fraction of the sterile soil indi-cates the importance of microbial activity oninteractions between humic acid and inositol andits metabolites.

The water-extractable fraction of the soilincubated with 14C(U)-myo-inositol was submittedto descending paper chromatography on large(56 × 23 cm) chromatogram sheets, as describedpreviously, and the paper chromatogram sub-jected to autoradiography to visualize the loca-tion of the separated 14C-labelled compounds


Table 4.1. Daily evolution of 14C carbon dioxide from an Andisol sandy loam treated with uniformlylabelled 14C myo-inositol, 14C phytic acid or 14C iron(III)-phytate. (From L’Annunziata and Gonzalez, 1977.)

14C-myo-inositol 14C-myo-inositol(non-sterile) (sterile) 14C-phytic acid 14C-iron(III) phytate

14C-CO2 Total CO214C-CO2 Total CO2

14C-CO2 Total CO214C-CO2 Total CO2

Day (%)a (meq) (%)a (meq) (%)a (meq) (%)a (meq)

1 38.25 0.644 − − 0.99 0.716 − 0.5982 4.49 0.493 − − 0.17 0.480 − 0.4403 3.63 0.486 − − 0.14 0.489 − 0.4294 2.80 0.460 − − 0.12 0.472 − 0.4475 2.17 0.414 − − 0.09 0.453 − 0.3976 1.77 0.424 − − 0.07 0.375 − 0.3707 1.67 0.367 − − 0.07 0.379 − 0.3748 1.32 0.384 − − 0.05 0.292 − 0.3179 1.44 0.424 − − 0.05 0.318 − 0.316

10 1.45 0.454 − − 0.06 0.382 − 0.36811 0.98 0.325 − − 0.04 0.286 − 0.26812 0.99 0.504 − − 0.05 0.350 − 0.314Total 61.0 ± 5.38 ± − − 1.90 ± 4.99 ± − 4.64 ±

0.44%b 0.42%b 1.4%b 0.14%b 2.6%b

aPercentage of radiolabel applied to the soil.bVariation between duplicate measurements or the average deviation expressed as a percentage of the mean.

Table 4.2. Phosphorus and radiocarbon contents in various fractions of soil incubated with uniformlylabelled 14C-myo-inositol under sterile and non-sterile conditions. (From L’Annunziata and Gonzalez, 1977.)

Phosphorusa (ppm) 14C-labelb (%)

Fraction Organic Inorganic Non-sterile Sterile

Soil-bound − − 31.54 0.42H2O-extractable − − 1.36 85.83HCl-extractable − Trace 4.54 13.90Humic acid −c 8.32 × 105 22.00 0.05Phosphates E 0.50 × 104 1.04 × 104 5.54 0.15Phosphates F 3.69 × 103 4.56 × 103 1.72 0.03Phosphates G 8.12 × 103 1.08 × 103 0.54 0.12Residue −c −c 36.92 1.01Total 1.68 × 104 ± 3.1%d 8.48 × 105 ± 4.1%d 104.16 ± 2.5%d 101.51 ± 2.6%d

aAnalysis of the non-sterile soil.bPercentage of total radiolabel remaining in the soil.cUndetermined because of high salt concentrations or interfering ions.dVariations between duplicate measurements or the average deviation expressed as a percentage of the mean.

(L’Annunziata et al., 1977). A typical autoradi-ogram is illustrated in Fig. 4.10.

Bands a1 and a2 of the water-extractable 14Cof the sterile and non-sterile soil were demon-strated to be due to the original unmetabolized14C-myo-inositol. It is of much interest to note thatthe water-extractable fraction of the sterile soilcontained only the original unmodified 14C-myo-inositol. The second band (b1) in Fig. 4.10 wasidentified to be 14C-chiro-inositol by infrared spec-trometry and thin-layer co-chromatography withstandards (Fig. 4.11). The optical activity of thechiro-inositol was not measured, but it wasassumed to be the D-chiro-enantiomer as L-chiro-inositol has not been identified in soils. On the

basis of the soil incubation studies with 14C(U)-myo-inositol, the carbon and phosphorus path-ways involving myo-inositol and its phosphateswere proposed (Fig. 4.12).

Recommendations for FutureResearch with Stable and

Radioactive Isotopes

Studies with inositol and its phosphates labelledwith radioactive isotopes, including 3H, 14C, 33Pand 32P as single or dual isotope labels, offer theadvantages of fast and easy real-time detection and


O

(a) (b)

O

a2

a1

b1

c1

d1

Fig. 4.10. Contact photographic prints made from X-ray film autoradiographs of paper chromatogramsof (a) the water-extractable fraction of the non-sterile soil incubated with 14C(U)-myo-inositol and (b) thewater-extractable fraction of the sterile soil incubated with 14C(U)-myo-inositol. (From L’Annunziata et al.,1977.) The chromatogram of (a) the non-sterile soil extract exhibited four bands marked a1, b1, c1 andd1, while the chromatogram of (b) the sterile soil extract exhibited only one band, marked a2. Bands a1and a2 were demonstrated to be the original 14C-myo-inositol, while band b1 was demonstrated to be14C-chiro-inositol, a microbial metabolite of 14C-myo-inositol. Bands c1 (weak) and d1 were not identified.

measurement of the isotope label. Flow-scintillationanalysis of liquid chromatography effluentsin homogeneous or heterogeneous flow cells pro-vides efficient analysis of beta-particle-emittingradionuclide tracers (L’Annunziata, 2003). High-performance liquid chromatography (HPLC)would be suitable mostly for the separation of thefree non-phosphorylated inositol stereoisomers andtheir related compounds (e.g. non-phosphorylatedinosose), such as those discussed in this chapter,which were subjected to acid hydrolysis to removethe phosphate groups prior to their chromato-graphic separation and structural elucidation.

Other separation techniques such as ion-exchangechromatography are currently the most efficient inseparating the inositol stereoisomers in phosphory-lated forms (see also Cooper et al., Chapter 3, thisvolume). The mobile phase in ion-exchange chro-matographic separation systems may containstrong chemical components that could precludethe use of on-line mass spectrometry or NMRspectrometry. In such cases fraction collection ofthe separated components as determined by theflow scintillation detector would be required priorto the preparation of samples for mass spectrome-try and NMR spectroscopy.

Whenever on-line spectroscopic techniquesare permitted the effluent from the flow scintilla-tion analyser can be connected with an ultravio-let absorbance detector, mass spectrometer orNMR spectrometer for molecular structureanalysis using chromatograph effluent splitting(Fig. 4.13). Such methods, often referred to asHPLC–FSA–UV–NMR–MS analysis systems,are described in detail elsewhere (L’Annunziata,2003). The free inositols would not yield anyultraviolet absorption, so the ultraviolet detectorwould be superfluous in such cases; however, theexpected inositol inosose (ketone) intermediates inthe chromatograph effluent would likely produceultraviolet absorption. Radiolabelled compoundsin the effluent would be detected by flow scintilla-tion prior to on-line mass spectrometry, NMRspectroscopy or fraction collection for off-line spectroscopy. 1H, 31P or natural-abundance13C NMR may be used for molecular structureelucidation. The NMR spectra on-line areobtained using a stop-flow method with resonancesignal acquisitions varying from several minutes tohours (Hansen et al., 1999; Smith et al., 1999;Bailey et al., 2000; Sweeney et al., 2000). The tech-niques are reviewed by L’Annunziata (2003).

Both homogeneous and heterogeneous flowscintillation analyses of the column chromato-graph effluents are possible. In homogeneousflow scintillation analysis a liquid scintillator isused to detect and measure the amount ofradioactivity in the chromatograph effluent. Insuch a case the flow scintillation analyser isequipped with a splitter to permit a fraction ofthe effluent to go on to the mass spectrometer,NMR spectrometer or fraction collector. If ion-exchange chromatographic separations of inositolphosphates do not permit on-line mass spectrom-etry or NMR due to strong chemical mobile


HO

HO

HOOH

30

cpm

20

SFO

10

OH

OHOH

HO

OH

OH

OH

OH

[14C]-myo-inositol

[14C]-chiro-inositol

Fig. 4.11. A thin-layer chromatogram (TLC) of theeluted paper chromatographic 14C-bands a1 andb1 of the non-sterile soil extract with authenticmyo- and chiro-inositols. (From L’Annunziata et al.,1977.) A recording of the 14C radioactivity (500counts per minute full scale) originating fromcomponents applied to the origin on the TLC plateis positioned directly above the plate. The lettersO and SF mark the origin and solvent front,respectively. The radioactivity of the chiro-inositolmetabolite represented ~4% of that of its myo-inositol precursor.

phases, the effluent splitter can be used to directa portion of the effluent to a fraction collector.The other fraction of the split effluent is mixed inliquid scintillator, radioactivity peaks plotted andcount rates (counts per minute) converted to dis-integration rates (disintegrations per minute) withprior radionuclide detection efficiency measure-ments. Reported detection efficiencies forradionuclide tracers in column chromatographeffluents by the homogeneous flow scintillationmethod are 20–60% for 3H, 70–95% for 14C,70–95% for 33P and 85–95% for 32P.

The heterogeneous flow scintillation methoddoes not require a liquid scintillator because it usesa solid scintillator (e.g. SolarScint®, Trademark ofPerkinElmer Life and Analytical Sciences) packedinto fine Teflon tubing through which thechromatograph effluent flows. This means thatheterogeneous flow scintillation analysis is less

cumbersome, less expensive and no chromato-graph effluent splitting is required for radioactivitymeasurement. Again, a fraction collector could beused whenever off-line mass spectrometry andNMR spectrometry are desirable. Heterogeneousflow scintillation detection efficiencies are some-what lower than the homogeneous method, being3% for 3H and 30% for 14C. Much higher detec-tion efficiencies would be expected for 32P, as thisradionuclide emits a beta particle with maximumenergy (Emax = 1710 keV), which is more than ten-fold greater than that of 14C (Emax = 155 keV).

Following are some of the questions thatmay be answered with radionuclide tracers:

1. Does the soil microbial epimerization of myo-inositol also yield scyllo-inositol and neo-inositol; ifso, does the epimerization occur via a mecha-nism similar to the epimerization of myo-inositolto chiro-inositol?


(b)

H

H

1

2

H

H

(c)

(d)

(d)

Ringcleavage

Othercatabolicreactions

CO2

HydralaseBerman and Magasanik(1966b)

H2O

DehydrataseBerman and Magasanik(1966a)

(a)(e)

Inositol phosphateisomers (e)

H

OH

OH

OH

OH

1H H

H

H

H

O

H

H

O

H

H

H

H

OH OH

OHO

H

OHHOHO

OH OH

H

HH

2HOHO

Fe+3

AI+3

Soilphosphate

(D)-chiro-Inositol(1,2,4/3,5,6)-inositol

Iron and aluminiumphytate (in acid soil)

Soilphosphate

Epimerization

Berman and Magasanik(1966a) and Posternak(1962)

Humic acid

myo -Inositol(1,2,3,5/4,6)-Inositol

Humic acid−inositol carboncomplexes or bonding

OH

H

H

OH

OH

OHHO

HO

Clay−inositol carbonabsorption Clay minerals

Acetobacter suboxidans

Fig. 4.12. Proposed carbon and phosphorus pathways of soil inositol including (a) the microbialepimerization of myo-inositol to (D)-chiro-inositol in soil. (From L’Annunziata and Gonzalez 1977.) Theepimerization of myo-inositol to chiro-inositol was demonstrated by the use of 14C-labelling; however, theoptical rotation of the chiro-inositol was not measured, hence the parenthesis around the dextrorotatorynotation. Other pathways of myo-inositol carbon in soils illustrated include (b) clay mineral absorption;(c) humic acid formation; (d) oxidation to carbon dioxide; and (e) phosphorus fixation throughphosphorylation and formation of iron and aluminium phytate complexes in acid soil.

2. What are the intermediates of the soil micro-bial epimerization of myo-inositol to chiro-inositol?3. Do soil myo-inositol phosphates also undergoepimerization?4. How stable are the alkaline earth (barium,calcium) salts or iron and aluminium salts of soilinositol phosphate stereoisomers as a function ofsoil pH and other factors, as the inositol phos-phate salts, under certain soil conditions, maycomprise a large pool of soil organic phosphorusnot readily available to plants or soil microor-ganisms?5. What are the rates of phosphorylation anddephosphorylation of inositol using 14C/32P dual-isotope labels?

The first three questions can be investigated withthe 14C isotope followed by mass spectrometryand 1H, 13C or 31P NMR. The flow scintillationanalyser, which may be coupled to an ion-exchange chromatograph column, is equipped todiscriminate and measure dual-isotope labelssuch as 3H–14C, 14C–32P and 33P–32P. The meas-

urement of single and dual radioisotope-labelledcompounds in column chromatograph effluents isreviewed by L’Annunziata (2003).

Examples of the possible soil microbialepimerization pathways (some yet unknown) ofmyo-inositol that may be resolved with the use of 3Hor 14C isotope tracer techniques are illustrated inFig. 4.14. Dual-labelled precursors such asuniformly ring-labelled 3H–14C(U)-myo-inositoland/or its phosphates together with the measure-ment of the 3H/14C isotope ratios of the myo-inositol precursor and its product compound(s) canprovide information on whether one or moreatoms of 3H are lost in the process. This wouldprovide insight into the mechanisms of myo-inositoltransformations in soil. The following hypotheticalcase may be taken as an example. If uniformlyring-labelled 3H–14C(U)-myo-inositol of known 3Hand 14C activities, measured in disintegrations perminute (DPM), is incubated in soil, and the 3H and14C activities of the product compound deter-mined, we can deduce whether one or more atomsof 3H are lost in the biochemical transformation.


Chromatography andNMR spectrometerconsole

FSA detectorandconsole

Splitter

MassspectrometerMass spectrometer

console

HPLC−NMR probe

Column

254UV detector

Injector

HPLCpump

NMRmagnet

Fig. 4.13. Instrumental set-up of the column chromatograph–ultraviolet–flow scintillation analysis–nuclearmagnetic resonance–mass spectrometry apparatus. Where ion-exchange liquid chromatographicseparations of inositol phosphates contain strong chemical mobile phases that do not permit on-linemass spectrometry or nuclear magnetic resonance (NMR) spectroscopy, the effluent splitter can be usedto direct a portion of the effluent on to a fraction collector. (Modified from Hansen et al., 1999.)


6

5

O

5

neo -Inositol(1,2,3/4,5,6)-Inositol

scyllo -Inositol(1,3,5/2,4,6)-Inositol

(D)-chiro-Inositol(1,2,4/3,5,6)-Inositol

4

* = 14C or 3H

3

21

6

4

3

2*

*

5

??

??

1 2

O

3

45

54

Phosphorylation

Inositol phosphate stereoisomers

3

21

6

6 *

* *

*

**

myo-Inositol(1,2,3,5/4,6)-Inositol

4

3

21

6

*

*

* *

*

1 2

3

4

O

(+)−Viboinososeintermediate

5

6M

icrobial epimerization in forest soil

* *

*

*

45

63

2 1

*

**

***

*

* *

*

* *

*

*

*

*

**

*

* *

*

*

*

*1

Fig. 4.14. The soil microbial epimerization of myo-inositol to (D)-chiro-inositol and proposed possibleepimerization pathways (marked with a question mark) of myo-inositol to the soil neo- and scyllo-inositols and their phosphates. The epimerization of myo-inositol to chiro-inositol was demonstrated bythe use of 14C-labelling. (From L’Annunziata et al., 1977.) The optical rotation of the chiro-inositol wasnot measured in this experiment with 14C, although the dextrorotatory character of the chiro-inositol waspreviously reported. (From Cosgrove, 1969; L’Annunziata, 1970; L’Annunziata et al., 1972.) The (+)-viboinosose intermediate illustrated above was identified to occur in Trifolium incarnatum by Scholdaet al. (1964). The inosose has not been demonstrated to occur in soil systems. The asterisks canrepresent either 3H or 14C isotope labels.

Table 4.3. Example of a hypothetical application of the 3H to 14C activity ratio measurements to the soilmicrobial transformation of myo-inositol.

DPMa Normalized DPM Normalized 3H/14C ratio

3H/14C activity ratio of uniformly ring-labelled 3H-14C(U)-myo-inositol precursor3H = 1200 (1200)(2.92) = 350014C = 3500 (3500)(1) = 3500 1:1 or 6:6

3H/14C activity ratio of metabolite (e.g. D-chiro-inositol)3H = 230 (230)(2.92) = 67214C = 800 (800)(1) = 800 672/800 or 5:6

aDPM = disintegrations per minute.

Table 4.3 provides hypothetical experimen-tal data to illustrate the application. The 3H and14C activities of the uniformly ring-labelled myo-inositol precursor are determined to be 1200 and3500 DPM, respectively. Because there are sixhydrogen and six carbon atoms on the inositolring, it is necessary to first normalize the 3H/14Cactivity ratio of the myo-inositol precursor so thatthe ratio equals 1:1 or 6:6. This is done, in thisexample, by multiplying the 3H activity by thefactor 2.92 necessary to bring up the 3H activityto equal that of the 14C and provide the requiredratio as illustrated in Table 4.3. If the metabolite,e.g. D-chiro-inositol, has 3H and 14C activities of230 and 800 DPM, respectively, we must firstmultiply the 3H activity by the same factor 2.92used to normalize the precursor activity ratio. Asillustrated in Table 4.3, this yields a 3H/14C ratioof 672 DPM/800 DPM or 5:6. From the changein the 3H/14C activity ratios from 6:6 to 5:6 wemay conclude the loss of one hydrogen atom inthe process and deduce an inosose intermediatesuch as that illustrated in Fig. 4.15. Activity ratiostudies with dual-labelled 14C/32P-myo-inositol

phosphates may also be carried out, the possibili-ties of which are limited only by the imaginationof the researcher. A review of the application ofdual-radioisotope activity ratios in numerousstudies of biochemical transformations is given byL’Annunziata (1984).

Studies with 3H, 14C, 33P or 32P as single- ordual-labelled myo-inositol or its phosphates offeradvantages of quick and facile tracing by flowscintillation analysis of soil microbial isotope-labelled metabolites in chromatograph effluents.The potential for the applications of isotopes inthe study of inositol phosphate transformations isindeed significant.

The work described here provides evidencefor two origins of the inositol stereoisomers andtheir phosphates in soils, namely plant residuesand epimerization of myo-inositol by soilmicrobes. Future studies with stable andradioactive isotopes can provide more informa-tion on the origins and transformations of theinositol stereoisomers and their role in thechemistry of soil phosphorus, soil fertility andsoil environment.


OH

OH

OH

OH

OH

OH

(+)-Viboinosose(loss of one 3H atom)

OHOH

OH

oOH

OH

myo-Inositol(1,2,3,5/4,6)-Inositol3H:14C or H :* = 6:6

1

2

**

**

*

*

H

HHH

H

H

H

H

H

H

H

H

H

H

H

H

OH

OH

OH

H

***

***

**

*

** *

OH

2

1

OH

D-chiro -Inositol(1,2,4/3,5,6)-Inostiol

3H:14C = 5:6

OH

Fig. 4.15. Epimerization of myo-inositol to D-chiro-inositol via (+)-viboinosose intermediate that involves theloss of a hydrogen atom from the inositol ring found in Trifolium incarnatum by Scholda et al. (1964). Theencircled ring hydrogen atoms and asterisks represent the molecular locations of 3H and 14C isotope labels.

References

Anderson, G. (1956) The identification and estimation of soil inositol phosphates. Journal of the Science of Food andAgriculture 7, 437–444.

Bailey, N.J.C., Cooper, P., Hadfield, S.T., Lenz, E.M., Lindon, J.C., Nicholson, J.K., Stanley, P.D., Wilson, I.D.,Wright, B. and Taylor, S.D. (2000) Application of directly coupled HPLC–NMR–MS/MS to the identifica-tion of metabolites of 5-trifluoromethylpyridone (2-hydroxy-5-trifluoromethylpyridine) in hydroponicallygrown plants. Journal of Agriculture and Food Chemistry 48, 42–46.

Bartha, R. and Pramer, D. (1965) Features of a flask and method for measuring the persistence and biologicaleffects of pesticides in soils. Soil Science 100, 68–70.

Berman, T. and Magasanik, B. (1966a) The pathway of myo-inositol degradation in Aerobacter aerogenes. Dehydrogenationand dehydration. Journal of Biological Chemistry 241, 800–806.

Berman, T. and Magasanik, B. (1966b) The pathway of myo-inositol degradation in Aerobacter aerogenes. Ring scission.Journal of Biological Chemistry 241, 807–813.

Beynon, J.H. and Williams, A.E. (1963) Mass and Abundance Tables for use in Mass Spectrometry. Elsevier, New York.Brownstein, S. (1959) Shifts in NMR absorption due to steric effects. II. Polysubstituted cyclohexanes. Journal of the

American Chemical Society 81, 1606–1608.Buchs, A., Charollais, E. and Posternak, T. (1968) Recherches dans la serie des cyclitols. XXXVII. Etude par spec-

trometrie de masse. Helvetica Chimica Acta 51, 695–707.Cosgrove, D.J. (1969) The chemical nature of soil organic phosphorus. II. Characterization of the supposed DL-

chiro-inositol hexaphosphate component of soil phytate as D-chiro-inositol hexaphosphate. Soil Biology andBiochemistry 1, 325–327.

Cosgrove, D.J. (1972) The origin of inositol polyphosphates in soil. Some model experiments in aqueous systemsinvolving the chemical phosphorylation of myo-inositol and the epimerization of myo-inositol pentaphosphates.Soil Biology and Biochemistry 4, 387–396.

Cosgrove, D.J. (1980) Inositol Phosphates: Their Chemistry, Biochemistry, and Physiology. Elsevier, Amsterdam, TheNetherlands.

Dyer, J.R. (1965) Applications of Absorption Spectroscopy of Organic Compounds. Prentice-Hall, Englewood Cliffs, NewJersey.

Greenland, D.J. (1956) The absorption of sugars by montmorillonite. I. X-ray studies. Journal of Soil Science 7,319–328.

Hansen, S.H., Jensen, A.G., Cornett, C., Bjørnsdottir, I., Taylor, S., Wright, B. and Wilson, I.D. (1999) High-per-formance liquid chromatography on-line coupled to high-field NMR and mass spectrometry for structure elu-cidation on constituents of Hypericum perforatum L. Analytical Chemistry 71, 5235–5241.

L’Annunziata, M.F. (1970) Soil–plant relationships and spectroscopic properties of inositol stereoisomers; the iden-tification of D-chiro- and muco-inositol in a desert soil–plant system. PhD thesis, University of Arizona, Tucson,Arizona. Available from University Microfilms International (UMI)/Bell and Howell Information andLearning (Dissertation Order No. 7104237), Ann Arbor, Michigan.

L’Annunziata, M.F. (1975) The origin and transformations of the soil inositol phosphate isomers. Soil Science Societyof America Proceedings 39, 377–379.

L’Annunziata, M.F. (1984) Agricultural biochemistry: reaction mechanisms and pathways in biosynthesis. In:L’Annunziata, M.F. and Legg, J.O. (eds) Isotopes and Radiation in Agricultural Sciences, Vol. 2: Animals, Plants, Foodand the Environment. Academic Press, London, pp. 105–182.

L’Annunziata, M.F. (2003) Flow scintillation analysis. In: L’Annunziata, M.F. (ed.) Handbook of Radioactivity Analysis,2nd edn. Elsevier, Amsterdam, The Netherlands, pp. 989–1062.

L’Annunziata, M.F. and Fuller, W.H. (1971a) Soil and plant relationships of inositol phosphate stereoisomers; theidentification of D-chiro- and muco-inositol phosphates in a desert soil and plant system. Soil Science Society ofAmerica Proceedings 35, 587–595.

L’Annunziata, M.F. and Fuller, W.H. (1971b) Nuclear magnetic resonance spectra of acetate derivatives of soil andplant inositol phosphates. Soil Science Society of America Proceedings 35, 655–658.

L’Annunziata, M.F. and Fuller, W.H. (1976) Evaluation of the mass spectral analysis of soil inositol, inositol phos-phates, and related compounds. Soil Science Society of America Journal 40, 672–678.

L’Annunziata, M.F. and Gonzalez, J. (1977) Soil metabolic transformations of carbon-14-myo-inositol, carbon-14-phytic acid and carbon-14-iron(III) phytate. In: Soil Organic Matter Studies, Vol. 1. International Atomic EnergyAgency, Publication No. IAEA-SM-211/66, Vienna, Austria, pp. 239–253.

L’Annunziata, M.F., Fuller, W.H. and Brantley, D.S. (1972) D-chiro-Inositol phosphate in a forest soil. Soil ScienceSociety of America Proceedings 36, 183–184.

L’Annunziata, M.F., Gonzalez, J. and Olivares, L.A. (1977) Microbial epimerization of myo-inositol to chiro-inositolin soil. Soil Science Society of America Journal 41, 733–736.

Lambert, J.B., Shurvell, H.F., Lightner, D.A. and Cooks, R.G. (1998) Organic Structural Spectroscopy. Prentice-Hall,Upper Saddle River, New Jersey.

Nakajima, M., Tomida, I., Kurihara, N. and Takei, S. (1959) Zur Chemie des Benzolglykols. V. Eine neueSynthesis der Inositole. Chemische Berichte 92, 173–178.

Peterbauer, T. and Richter, A. (1998) Galactosylinositol and stachyose synthesis in seeds of Adzuki bean. PlantPhysiology 117, 165–172.

Posternak, T. (1962) Scyllo-Inosose (myo-inosose-2): Bacterial oxidation of myo-inositol. In: Whistler, R.L.,Wolfram, M.L., Bemiller, J.N. and Shafizadeh, F. (eds) Methods in Carbohydrate Chemistry. Academic Press, NewYork, p. 294.


Reed, R.I., Reid, W.K. and Wilson, J.M. (1962) The mass spectra of some flavones and carbohydrates. In: Advancesin Mass Spectrometry, Vol. 2. Pergamon Press, New York, pp. 416–427.

Richter, A. Thonke, B. and Popp, M. (1990) D-1-O-methyl-muco-Inositol in Vicum album and members of theRhizophoraceae. Phytochemistry 29, 1785–1786.

Scholda, R., Billek, G. and Hoffmann-Ostenhof, O. (1964) Biosynthesis of cyclitols. VIII. Mechanism of theconversion of meso-inositol to D-pinitol and D-inositol in Trifolium incarnatum. Monatshefte für Chemie 95,1311–1317.

Smith, R.M., Chienthavorn, O., Wilson, I.D., Wright, B. and Taylor, S.D. (1999) Superheated heavy water as theeffluent for HPLC–NMR and HPLC–NMR–MS of model drugs. Analytical Chemistry 71, 4493–4497.

Sweeney, D.J., Lynch, G., Bidgood, A.M., Lew, W., Wang, K.-Y. and Cundy, K.C. (2000) Metabolism of theinfluenza neuraminidase inhibitor prodrug oseltamivir in the rat. Drug Metabolism and Disposition 28, 737–741.

Turner, B.L., Papházy, M.J., Haygarth, P.M. and McKelvie, I.D. (2002) Inositol phosphates in the environment.Philosophical Transactions of the Royal Society, London, Series B 357, 449–469.

Wanek, W. and Richter, A. (1995) Purification and characterization of myo-inositol 6-O-methyltransferase fromVigna umbellate Ohwi et Ohashi. Planta 197, 427–434.


5 Isolation and Assessment ofMicroorganisms That Utilize Phytate

Jane E. Hill1 and Alan E. Richardson2

1Environmental Engineering Program, Yale University, 9 Hillhouse Avenue, PO Box8286, New Haven, CT 06520, USA; 2CSIRO, Plant Industry, PO Box 1600,

Canberra, ACT 2601, Australia


The first phytase was discovered in the earlierpart of the last century (Suzuki and Takaishi,1907; Dox and Golden, 1911), but it was not untilthe efforts of Cosgrove and others in the 1960s(Cosgrove, 1980) that interest in identifying, char-acterizing and commercializing phytate-degradingorganisms and their enzymes intensified. Indeed,it is becoming increasingly apparent that a rangeof microorganisms possess the ability to utilizephytate in their environment (Shieh and Ware,1968; Irving and Cosgrove, 1971; Power andJagannathan, 1982; Richardson and Hadobas,1997; Yanke et al., 1998). These microorganismsoccur in a variety of diverse habitats and do notemploy an identical method to degrade phytate.Phytases, defined as enzymes that initiate thecleavage of myo-inositol hexakisphosphate, arenow classified into four groups based on commonbiochemical and catalytic mechanisms. These are:(i) histidine acid phosphatases, which are wide-spread in fungi (Ullah et al., 1991; Van Etten et al.,1991); (ii) cysteine phosphatases, which have beenidentified in rumen bacteria (Yanke et al., 1999;Chu et al., 2004); (iii) β-propeller phytases, whichoccur mainly in the bacilli bacteria (Kim et al.,1998b; Ha et al., 1999; Shin et al., 2001; Oh et al.,2004); and (iv) purple acid phosphatases, whichoccur in a variety of organisms. The phytases aredescribed in detail elsewhere in this volume (seeMullaney and Ullah, Chapter 7).

The existence of phytate-degrading micro-organisms challenges the conventional perception

that inositol phosphates are recalcitrant in theenvironment and of limited biological availability.This has arisen from the strong stabilization ofinositol phosphates following sorption to clays orprecipitation with metals (see Celi and Barberis,Chapter 13, this volume), their abundance in soils(see Turner, Chapter 12, this volume) and theirlimited availability to plants (see Richardson et al.,Chapter 15, this volume). Our understanding ofphytate-degrading microorganisms has been limi-ted by a lack of ecological information and thedifficulty in isolating and assessing the causativemicroorganisms in vivo. This chapter describesprocedures for screening phytate-degrading micro-organisms and characterizing their phytaseactivity, and provides recommendations for futurestudies. The major groups of microorganisms thathave been isolated and characterized are summar-ized and current ecological information for theseorganisms is discussed.

Assessing Microorganisms forPhytate-degrading Activity

Microorganisms able to degrade phytate havebeen isolated from a range of terrestrial andaquatic environments and their optimal cultiva-tion conditions are often genera-specific. Theyinclude yeasts, filamentous fungi and Gram-positive as well as Gram-negative bacteria. Three

microorganisms, in particular Aspergillus niger,Escherichia coli and Peniophora lycii, have been stud-ied in detail and are produced commercially asan additive for animal feeds (Simon and Igbasan,2002). However, the inherent variety in phytate-degrading microorganisms and the phytases thatthey produce suggests that other types of organ-isms and phytases await characterization. Thediversity of phytate-degrading microorganismsis immense and requires a more detailedcategorization than that used for the enzymesthemselves. Detailed information on phytate-degrading microorganisms and the ecosystems inwhich they are found is fundamental to improv-ing our understanding of inositol phosphates inthe environment.

Screening for phytate utilization

The most common procedure for screeningmicroorganisms for potential to degrade phytateinvolves growth on solid laboratory media con-taining sodium- or calcium-phytate (Shieh andWare, 1968; Cosgrove et al., 1970; Richardsonand Hadobas, 1997). Calcium-phytate forms aprecipitate at the pH range used in most media(pH 5–8) and can indicate the presence of phy-tate-degrading organisms in two ways. First, amicroorganism growing on phytate as the solesource of phosphorus should only be able to do so

due to its ability to release an extracellular phy-tase or by uptake through the outer membraneand its subsequent dephosphorylation within theperiplasm (see Greiner, Chapter 6, this volume).Second, a clear zone around colonies that growon calcium-phytate media indicates capacity tosolubilize phytate presumably through the pro-duction of acids or chelates (Richardson andHadobas, 1997; Mukesh et al., 2004; Fig. 5.1).Appropriate controls must be included to showthat isolates do not grow when presented with amedium with all the ingredients except phytate.False positives may result from trace amounts ofnon-phytate phosphorus being present in theinocula or medium ingredients. A diagnosticallyfalse zone of clearing around a colony may becaused by the production of acid alone.

Not all organisms that grow on solid mediawill grow easily in liquid culture (Richardson andHadobas, 1997), so isolates can also be assessedfor phytate utilization using liquid media.Growth of an organism in liquid culture contain-ing phytate as the sole source of phosphorusshould, with correct controls, provide a betterindication of the ability of an organism todegrade phytate. Care must be taken to ascertainthat the reason for growth is the degradationof phytate. Adequate controls showing a lack ofgrowth as well as the removal of trace amounts ofinorganic phosphate from the inocula are essen-tial. Even with these precautions, growth maystill occur (e.g. from phosphorus derived from

62 J.E. Hill and A.E. Richardson

(a)

Sodium-phytate Calcium-phytate

pH 7

pH 5

0Bac

teria

l gro

wth

(lo

g 10

cells

/ml)

Sod

ium

-phy

tate

con

tent

(m

g/m

l)

Pho

spha

te r

elea

sed

(% to

tal)

2.0

4.0

6.0

8.0

0

20

40

60

80

0

2

4

6

8

10

12 24 36 48

Time (h)

10.0

()

()

()

(b)

Fig. 5.1. Screening for phytate-degrading microorganisms. (a) Growth of Pseudomonas sp. strainCCAR59 on agar containing sodium-phytate and calcium-phytate as the sole source of carbon andphosphorus at pH 7 and 5. (b) Degradation of sodium-phytate and release of phosphate to the culturesupernatant during growth (pH 5.5) of strain CCAR59 in liquid culture. (From Richardson and Hadobas,1997.)

lysis of the inoculum cells). Such procedures,therefore, only indicate the potential to degradephytate and should not be used in isolation. Bothsolid and liquid screening procedures do not con-firm phytase activity, and characterization ofphytase activity and/or enzyme purification isnecessary, as outlined below.

It should also be considered whether micro-organisms are using phytate as a source of carbon,phosphorus or both. While some cultured organ-isms can survive on phytate as a sole source of car-bon and phosphorus (e.g. Pseudomonas spp. andKlebsiella aerogenes; Irving and Cosgrove, 1971;Tambe et al., 1994), other organisms may not(Richardson and Hadobas, 1997). A further import-ant point is the nature of the carbon source used.Phosphorus bioavailability is often dependent onthe ability of an organism to solubilize the source ofphosphorus. Substrates such as citrate, which is achelating agent, may make phytate more bioavail-able by influencing substrate solubility (Hernandezet al., 2003; Munoz and Valiente, 2005).

Case study of the isolation of phytate-degrading Pseudomonas spp. from soil

Two strategies to isolate phytate-degradingmicroorganisms were used by Richardson andHadobas (1997). In one approach, agar mediumcontaining sodium-phytate was used to identifymicroorganisms with the potential to degradephytate. This indicated that 63% of isolates wereable to grow on the defined medium containingphytate as the sole source of phosphorus and car-bon, relative to tryptic-soy agar (a general growthmedium for heterotrophic bacteria). When a 100-member subset of these microorganisms wasgrown in liquid culture of the same composition,none were able to grow. However, 39% of theisolates were able to grow when citrate wasadded to the medium, although none were ableto liberate phosphate from phytate as determinedby the presence of molybdate-reactive phosphatein the culture supernatant.

The second approach used enrichment cul-ture and was more successful in identifying iso-lates that could degrade phytate. Liquid culturescontained phytate as the sole source of phospho-rus either with or without the addition of citrateas a carbon source (Richardson and Hadobas,

1997). After purification of broth isolates by agarmedium of the same composition, 44% of isolateswere able to grow when reinoculated into theoriginal broth medium. Of these, 4% liberatedphosphate in the culture supernatant, from whichsix genetically unique bacterial strains were iso-lated. These isolates showed greatest similarity toPseudomonas spp., which are a ubiquitous group ofmicroorganisms found in soil. Growth of oneof these isolates (strain CCAR59) on sodium-phytate and calcium-phytate media as the solesource of carbon and phosphorus is shown inFig. 5.1, with solubilization of calcium-phytatebeing evident at pH 5. This example highlightsthe importance of using a combination of growthconditions and assay procedures to verify thatmicroorganisms are able to utilize phytate.

Characterization of phytase activity

Following the isolation of microorganisms thathave the ability to degrade phytate in laboratorymedia it is necessary to confirm phytase activity.Ideally this is achieved through purification of theenzyme and characterization of its kinetics,although this is often a challenging and time-consuming task. First, organisms must be grownin culture to generate a sufficient amount ofenzyme to enable purification. Then, a series ofpurification steps, such as ammonium sulphatefractionation and chromatographic separation,are used to purify the enzyme. Molecular weight,isoelectric point, specific activity, thermal stabilityand pH optima can then be evaluated (e.g.Greiner et al., 1993; Shimizu, 1993; Greiner et al.,1997; Choi et al., 2001).

Colorimetric assays, which measure freephosphate in solution, are useful for assessingphytase activity. Typically, microbial extracts orextracellular components are incubated in thepresence of sodium-phytate under defined condi-tions (i.e. temperature, pH, etc.) and releasedphosphate is determined at intervals. Thisapproach is primarily used for initial assessmentof phytase activity from different microorganisms,subsequent tracking of enzymes during purifica-tion procedures, determination of kinetics andestablishment of an unequivocal link betweenbiological synthesis of the enzyme and the utiliza-tion of phytate.

Microorganisms That Utilize Phytate 63

Antibody-based technologies have also beenused to track purification as well as to localizephytase in microorganisms and tissues (Ullah andGibson, 1987; Golovan et al., 2001). Immuno-assays use antibodies to recognize highly specificmotifs (generally protein domains) that are gener-ated from the immune response of animals.Mono- and polyclonal antibodies have beenraised for phytases from A. niger (Ullah andGibson, 1987; Hostetler et al., 2005) and E. coli(Golovan et al., 2001).

Case study of phytase purification andcharacterization from Klebsiella terrigena

Klebsiella terrigena is a soil bacterium that can utilizephytate as the sole source of phosphorus. Phytaseproduction is greatest when the culture reaches thestationary phase and the activity of the enzyme isstimulated by the presence of phytate (see Greiner,Chapter 6, this volume). Greiner et al. (1997) puri-fied the enzyme, characterized its substrate speci-ficity and kinetics, and determined the products ofenzymatic hydrolysis using high-performance liq-uid chromatography (HPLC). Protein purificationto this extent (i.e. as outlined in Table 5.1) is nec-essary to characterize the enzyme. Preparationpurity is ascertained using sodium dodecyl sulph-fate–polyacrylamide gel electrophoresis (SDS–PAGE), which can also be used to indicate protein

molecular mass and the extent of glycosylation.Once purified, phytases from different sources canbe compared (Table 5.2).

Assessment of phytase activity byanalytical methods

Major techniques that have been applied forassessing the catalytic function of phytases fromdifferent microorganisms include ion chro-matography, HPLC and 31P nuclear magneticresonance (NMR) spectroscopy. These tech-niques are particularly useful for investigatingthe presence of lower-order inositol phosphatesand inorganic phosphates to establish pathwaysof degradation (Freund et al., 1992; Skoglundet al., 1997; Cade-Menun, 2005). For example,Greiner et al. (1997) used ion pair chromatog-raphy to conduct a time course study of thedegradation of sodium-phytate by K. terrigenaphytase (Fig. 5.2). The results showed the accu-mulation of an inositol trisphosphate end prod-uct, although the presence of positional isomerscould not be determined.

The dephosphorylation pathway for severalBacillus spp. that degrade phytate has similarlybeen determined (Greiner et al., 2002). In thiscase, the genes responsible for phytate degrad-ation from Bacillus subtilis 168, B. amyloliquefaciensATCC 15841 and B. amyloliquefaciens 45 were


Table 5.1. Purification of phytase from Klebsiella terrigena showing higher specific activity withincreasing purity. (From Greiner et al., 1997.)

Purification step Total activitya (U) Specific activityb (U/mg) Recoveryc (%)

Crude extract 268.0 0.5 –(NH4)2SO4 precipitation 255.0 1.5 95CM-Sepharosed 178.0 55.6 66DEAE-Sepharosee 112.0 80.0 42Mono Sf 90.0 102.3 34Sephacryl S-200g 76.0 205.0 28

aTotal activity (U) is determined where 1 unit liberates 1 µmol phosphate per minute.bSpecific activity is the total activity divided by total protein in the sample.cRecovery is the total activity in the step divided by the total activity of the crude extract expressed as a percentage.dCM-Sepharose is the cationic exchange resin carboxymethyl-Sepharose.eDEAE-Sepharose is the anionic exchange resin diethylaminoethyl-Sepharose.fMono S is a cation exchange resin.gSephacryl S-200 is a hydrophilic resin.

cloned and expressed in a common strain ofbacilli (B. subtilis MU331), and were comparedto phytases from E. coli, A. niger and rye (Secalecereale L.). Degradation products were determinedby HPLC and demonstrated that Bacillus spp.had two independent pathways for degradationof phytate. This initially led to an accumulationof either D/L-myo-inositol 2,4,5-trisphosphate ormyo-inositol 2,4,6-trisphosphate and subsequentlyto the generation of myo-inositol 2-monophos-phate (for further details see Greiner, Chapter 6,this volume).

Sources of Phytase and Expressionin Microorganisms

Phytate-degrading microorganisms

Phytate-degrading microorganisms have been iso-lated from a wide range of environments, includ-ing marine and fresh water ecosystems, soils,sediments and the gastrointestinal tract of animals(Table 5.3). Although no systematic study hasbeen conducted across different environments, itis evident that the phytase phenotype is manifestin a variety of habitats. This includes a variety ofsoils (Richardson and Hadobas, 1997), organic-rich and organic-poor locations (Choi et al., 2001;Kim et al., 2003) and anaerobic environments(Yanke et al., 1999). The anaerobic environment isof particular interest, because inositol phosphatesappear to be degraded rapidly under anaerobicconditions (Suzumura and Kamatani, 1995a) andare absent from submerged wetland soils whereanaerobicity is common (Turner and Newman,2005). Anaerobic environments may therefore beimportant in the inositol phosphate cycle andan important source of novel phytate-degradingmicroorganisms.

The mixture of intracellular and extracellu-lar enzymes (Table 5.3) is also significant andsuggests that microorganisms use different mech-anisms to hydrolyse inositol phosphates, eitherin the external environment or within theperiplasm. Extracellular enzymes are generallymore tolerant to pH and temperature fluctuationand show greater resistance to proteolytic degrad-ation (see George et al., Chapter 14, this volume).These traits will not only affect the persistenceand effectiveness of phytase from differentmicroorganisms in different environments, butare also attractive features for the commercialdevelopment of phytases.

Production of microbial phytases

Both native and recombinant phytase enzymeshave been assessed in an effort to optimize pro-duction. Filamentous fungi in particular havebeen utilized extensively for their high pro-duction efficacy. However, E. coli and yeast(Saccharomyces cerevisiae) (Phillippy and Mullaney,


Table 5.2. Kinetics constants (kcat) for degradationof myo-inositol hexakisphosphate and its lower-order esters by phytase from Bacillusamyloliquefaciens (strain ATCC 15841). (FromGreiner et al., 2002.)

Substrate kcat (/s)

myo-Inositol hexakisphosphate 18.5ca

D-myo-Inositol 1,2,4,5,6-pentakisphosphate 16.0a

D-myo-Inositol 1,2,3,5,6-pentakisphosphate 7.3b

D-myo-Inositol 1,2,3,4,5-pentakisphosphate 7.5b

D-myo-Inositol 1,2,5,6-tetrakisphosphate 10.1d

D-myo-Inositol 1,2,6-trisphosphate 1.9emyo-Inositol 2-monophosphate 0.12g

aMeans accompanied by a different letter are significantlydifferent (P < 0.05).

0

10

20

30

40

0 120 240 360 480

Time (min)

Inos

itol p

hosp

hate

s (µ

g)

IP6

IP5 IP4

IP3

Fig. 5.2. Time course for the degradation ofsodium-phytate by Klebsiella terrigena phytase asdetermined by ion pair chromatography. (FromGreiner et al., 1997 with permission fromElsevier.) myo-Inositol hexakisphosphate (IP6) ishydrolysed to a myo-inositol trisphosphate (IP3)end product. IP4, myo-inositol tetrakisphosphate;IP5, myo-inositol pentakisphosphate.

1997; Han et al., 1999; Kerovuo et al., 2000;Sajidan et al., 2004) and plant systems have alsobeen employed (Pen et al., 1993; Li et al., 1997;Kusnaki et al., 1998; Ponstein et al., 2002). Thereis still much to be understood about the basicstimuli and repression systems that are associatedwith the expression of phytases in native systems.However, phosphorus limitation, in the form ofeither a low intracellular or ambient phosphorusconcentration, is a key stimulus for phytase pro-duction in many microorganisms. This impliesthat phosphorus limitation is paramount to theexpression of genes for phytase synthesis inmicroorganisms. However, there are notableexceptions to this: e.g. the rumen bacteriareported by Yanke et al. (1998), which live in anenvironment rich in phosphorus.

Optimization of cultures for expression of phytase

Growth conditions significantly affect the expres-sion and production of phytase, but no genericmethodology exists to assess this and few trendsare apparent. E. coli has two periplasmic phos-phatases that can accept phytate as a substrate.The first, agp, has a broad substrate range, isconstitutively expressed and may be used to scav-enge glucose from glucose 1-phosphate (Wanner,1990). The second, appA, has a narrow substraterange and its expression (and production) is stimu-lated by entry into the stationary phase of growth,a low ambient phosphate concentration andanaerobic conditions (Touati et al., 1987; Greineret al., 1993; see Greiner, Chapter 6, this volume).


Table 5.3. Source of phytate-degrading microorganisms, their environment of isolation, oxygenrequirement and location of enzyme expression.

Oxygen Enzyme Microorganism Environment requirementa locationb Reference

Bacillus sp. KHU-10 Boiled rice A EX Choi et al. (2001)Bacillus sp. DS11 Cattle shed floor A EX Kim et al. (1998a)Citrobacter braakii YH-15 Sea water A IN Kim et al. (2003)Enterobacter sp. 4 Soil of leguminous A EX Yoon et al. (1996)

plantsKlebsiella oxytoca MO-3 Soil A IN Jareonkitmongkol

et al. (1997)Lactobacillus sanfrancis- Italian sourdough A IN De Angelis et al.

censis CB1 (2003)Myceliophthora Cellulosic waste A EX Mitchell et al. (1997)

thermophila (48102)Penicillium simplicissimum Soil A EX Tseng et al. (2000)Pseudomonas syringae Cattle farm soil A IN Cho et al. (2003)

MOK1Pseudomonas spp. Soil A IN Richardson and

Hadobas (1997)Paramecium Laboratory stock A IN Freund et al. (1992)

tetraurelia 51s (fresh water organism)

Pichia anomala Dried flower buds A IN Vohra and Satyanarayana (2002)

Rhizopus oligosporus Tempehc A IN/EX Sutardi and Buckle CT11K2 (1988)

Selenomona ruminatium Rumen fluid AN EX Yanke et al. (1999)

aA/AN refers to aerobic or anaerobic production, respectively.bExpression is either intracellular (IN) or extracellular (EX).cA fermented food typically made from soybean that is popular in South-east Asia.

Induction of the PHO regulon is not implicated inthe synthesis of the enzyme (Touati et al., 1987)nor is it induced by phytate (Greiner et al., 1993).Under aerobic growth conditions, growth restric-tion by low inputs of carbon, nitrogen and sul-phur had no influence on the production of eitherof the E. coli enzymes (Touati et al., 1987).

Bacillus spp. (strain KHU-10) produce a highlevel of extracellular phytase in the stationaryphase when grown in complex media containingmaltose, peptone and beef extract (Choi et al.,1999). Stimulation of phytase production in B.subtilis has also been observed by the addition ofphytate to the cultivation medium (Kim et al.,1999b). In more simple but defined media,Bacillus phytases require calcium ions for effectivephytase activity. However, this requirementappears to influence only the stability of theenzyme rather than regulate its production(Shimizu, 1992; Choi et al., 1999). In more com-plex media (e.g. wheat bran medium), factorsthat contribute to high phytase production arenot well characterized, but their impact on pro-duction and activity of phytase is significant(Shimizu, 1992; Choi et al., 1999). The low solu-bility of phytate in wheat bran is thought to resultin a controlled release of substrate, which directlyregulates phosphate in the medium and, there-fore, enzyme production. However, other factorsin the medium may also be important.

Production of phytase in fungi is typicallyenhanced by low phosphate in the medium andis repressed at higher concentrations (Shieh andWare, 1968; Han and Gallagher, 1987; Vatsand Banerjee, 2002; Kim et al., 2003). Muta-genesis of cultures (e.g. through irradiation)results in increased phytase production: e.g. in A.niger NRRL3135 (Chelius and Wodzinski, 1994).However, a decrease in enzyme productionoccurs when the medium contains a high concen-tration of glucose and/or is poorly aerated.

Saccharomyces cerevisiae produces three extra-cellular phosphatases (PHO5, PHO10 andPHO11) that can hydrolyse phytate as well as arange of other phosphate monoesters (Nakamuraet al., 2000; Andlid et al., 2004). These genes areassociated with the PHO regulon and respond tolow levels of extracellular phosphate (Lemireet al., 1985) in a complex regulatory framework(Ogawa et al., 1993; Kaffman et al., 1994; seeGreiner, Chapter 6, this volume).

Production of phytases on a large scale hasalso been undertaken using submerged and solid-state fermentation technologies. As with lab-scalecultures, each strain has optimal conditions,including the rate of aeration, temperature, inocu-lum conditions and media components. Thesehave been best studied for A. niger NRRL3135(Howson and Davis, 1983; Han and Gallagher,1987; Ullah and Gibson, 1987; Ebune et al., 1995;Krishna and Nokes, 2001). Submerged fermenta-tion studies for E. coli and Bacillus sp. DS11 showthat phytase production was essentially the sameas for smaller-scale culture studies (Kim et al.,1998a; Kleist et al., 2003).

Large-scale production is an essential subse-quent step for the commercialization of enzymes.Generally this is best achieved when phytase-encoding genes are cloned and expressed inrecombinant microorganisms using expressionvectors, where protein production can more eas-ily be optimized and yields are often substantiallyhigher (Quax, 1997; Han et al., 1999; Kim et al.,1999a; Rodriguez et al., 2000a; Xiong et al., 2004;Table 5.4). In contrast, optimization of cultureconditions for the expression of ‘native’ enzymesis more difficult. Observations based on composi-tion of the media are often inconclusive, due inlarge part to the complexity of the regulation ofphytase gene expression and conditions forenzyme activation in diverse microorganisms.These difficulties notwithstanding, culture opti-mization can provide valuable insight into theregulation of the enzyme in vivo.

Properties of phytases frommicroorganisms

Catalytic and physicochemical properties of phy-tases from some representative microorganismsare shown in Table 5.5. Phytase from Citrobacterbraakii has the highest reported specific activity(Kim et al., 2003). The E. coli appA phytase withan optimal pH 2.5 has the highest reported kcat(Dassa et al., 1982; Greiner et al., 1993) and hasbeen subjected to a number of mutagenic modifi-cations (Golovan et al., 2000; Rodriguez et al.,2000b). Similarly, phytase (phyA) from A. nigerNRRL 3135 (formerly A. ficuum) has been exten-sively studied in terms of catalytic and physical


properties and mutational analysis (e.g. Wysset al., 1999a,b; Lehmann et al., 2002; Tomschyet al., 2002). A. niger, with two pH optima (2.2 and5.0–5.5; Ullah and Gibson, 1987), is a particu-larly useful enzyme due to its ability to functioneffectively under gastrointestinal conditions rele-vant for swine and poultry. The cloning andexpression of this phytase (Mullaney et al., 1991;van Hartingsveldt et al., 1993) has led to its com-mercial development as a feed additive (e.g.Natophos®). Likewise the phytases from P. lycii(Lassen et al., 2001) and E. coli (Fairley, 1998)have been developed commercially.

From a commercial perspective, phytasesdestined for addition to the diet of animals needto meet a number of specific requirements.Animal feeds are often produced through a pellet-ing process at temperatures between 65°C and80°C. Thermostability of the phytase is thereforedesirable. Phytases are also required to functionwithin the gastrointestinal tract of various animalsthat have unique body temperature and digestivepH (Riley and Austic, 1984; Radcliffe et al., 1998;

Ramseyer et al., 1999). For example, the adultbody temperature for swine is around 39°C,whereas it is between 5ºC and 18°C in fish. Age-dependent variations must also be considered andresistance of phytases to proteolytic cleavage isimportant (Wyss et al., 1999b). Modification of phy-tases to meet such requirements has been studiedextensively (e.g. Wyss et al., 1999a,b; Lehmannet al., 2002; Tomschy et al., 2002).

The Ecology of Phytate-degradingMicroorganisms

Inositol phosphates exist in a wide range of envi-ronments including arable, forest and grasslandsoils, as well as fresh water and marine sediments(Turner et al., 2002; see Turner, Chapter 12, andMcKelvie, Chapter 16, this volume). Given thatproductivity in many of these ecosystems can belimited by the availability of phosphorus, why doinositol phosphates persist when microorganismsare present that can potentially utilize them?


Table 5.4. Host expression systems and cultural conditions for expression of phytases from diversemicroorganisms.

Microorganism and Temperature source of phytase Expression host pH optima optima (ºC) Reference

Aspergillus niger E. coli 5.1 43–50 Phillippy and Mullaney (1997)

Saccharomyces 2.0–2.5, 5.0–5.5 55–60 Han et al. (1999)cerevisiae

Pichia pastoris 2.5, 5.5 60 Han and Lei (1999)A. niger (native) 2.2, 5.0–5.5 55–58 Ullah and Gibson

(1987), Wyss et al.(1999a,b)

Escherichia coli E. coli 4.5 55 Golovan et al. (2000)P. pastoris 3.5 60 Rodriguez et al.

(2000a)Pseudomonas 4.0 55 Dharmsthiti et al.

putida (2005)E. coli (native) 4.5 55–60 Greiner et al. (1993)

Bacillus subtilis 168 B. subtilis 7.0 55 Tye et al. (2002)B. subtilis 168 6.0–7.5 55–60 Shimizu (1992),

(native) Kerovuo et al. (1998)B. licheniformis B. subtilis 7.0 65 Tye et al. (2002)

B. licheniformis 6.0–7.5 55–60 Shimizu (1992), (native) Kerovuo et al. (1998)

Bacillus sp. DS11 E. coli – 70 Kim et al. (1998b)Bacillus sp. DS11 7.0 70 Kim et al. (1998a)

(native)

Microorganism

s That U

tilize Phytate

69

Table 5.5. Catalytic and physicochemical properties of phytases from diverse microorganisms (values denoted by a dash are not available).

Temperature Specific Substrate Microorganism pI pH optima optima (ºC) activitya (U/mg) km (µmol) range Reference

Aspergillus niger 4.5–5.2 2.5, 5.0–5.5 55–58 50–103 10–40 Narrow Ullah and Gibson (1987), NRRL3135 Wyss et al. (1999a)

A. terreus 5.0 5.0–5.5 70 142–196 11–23 Narrow Yamada et al. (1968), Wyss et al. (1999a,b)

Bacillus licheniformis 5.0 4.5–6.0 55–60 – – Narrow Tye et al. (2002)B. subtilis 6.3–6.5 6.0–7.5 55–60 9–15 50–500 Narrow Shimizu (1992), Kerovuo

et al. (1998)Bacillus sp. DS11 5.3 7.0 70 – 550 Narrow Kim et al. (1998a)Citrobacter braakii – 4.0 50 3457 460 Narrow Kim et al. (2003)Escherichia coli 6.0–7.4 4.5 55–60 811–1800 130–630 Narrow Greiner et al. (1993)Lactobacillus 5.0 4.0 45 6–73 – Middle De Angelis et al. (2003)

sanfranciscensisKlebsiella terrigena 5.0–6.0 55 205 300 Middle Greiner et al. (1997)Penicillium simplicissimum 5.8 4.0 55 3 – Broad Tseng et al. (2000)Peniophora lycii 3.6 4.0–4.5 45 – – Narrow Lassen et al. (2001)Pseudomonas syringae – 5.5 40 – 380 Narrow Cho et al. (2005)

MOK1Saccharomyces cerevisiae – 2.0–2.5, 5.0–5.5 55–60 – – Broad Han et al. (1999)Schwanniomyces castellii – 4.4 77 418 38 Broad Segueilha et al. (1992)

aDetermined at 37ºC, with the exception of Schwanniomyces castellii, which was determined at 70ºC.

Perhaps the most important factor that reg-ulates the availability of inositol phosphates tomicroorganisms is the formation of strong andstable complexes between inositol phosphatesand metals such as iron, aluminium, copper, cal-cium and magnesium (Vohra et al., 1965; see Celiand Barberis, Chapter 13, this volume). Persist-ence of inositol phosphate in soils therefore maybe linked to the ability of microorganisms to solu-bilize and access these forms of phosphorus.Alternatively, many soils do not represent phos-phorus-limited environments for microorganisms(Jakobsen et al., 2005), reducing the need formicroorganisms to utilize inositol phosphates. Inthis scenario, inositol phosphate concentrationswould be linked to the degree of phosphorus lim-itation, and there is tentative evidence for this insome soils (see Turner, Chapter 12, this volume).

To date much of our understanding of phy-tase from microorganisms has been driven by thepotential for commercial phytase production,rather than to obtain an ecological understand-ing. Clearly, there is a need to better understandthe ecological importance of microorganisms thathave the potential to degrade phytate in differentecosystems.

Function of phytases in differentecosystems

Plate and liquid culture screening assays are asimple and effective means to obtain generalinformation on potential phytate-degrading activ-ity of microorganisms in environmental samples.However, as described above, such assessmentmay result in erroneous estimates of the numberof organisms with phytate-degrading capability(Richardson and Hadobas, 1997; Fig. 5.1). Forexample, Greaves and Webley (1965) estimatedsoil and rhizosphere populations of phytate-degrading bacteria to constitute between 30%and 48% of the total population using plate-basedscreening methods. While this may be an overesti-mate, Unno et al. (2005) recently isolated andcharacterized a large number of microorganisms,predominantly Burkholderia spp., from the rhi-zosheath and rhizoplane of lupin (Lupinus albus L.)plants. Interestingly, many of these isolates werealso able to promote plant growth, suggesting theimportance of phytate hydrolysis in close proxim-ity to plant roots (see Richardson et al., Chapter

15, this volume). Although such screening proce-dures do not unequivocally demonstrate the func-tion of phytate-degrading microorganisms in soil,they do indicate a potential role. Further work isneeded to establish the function of phytate-degrading microorganisms in such environments.An important advance in this area is the develop-ment of a substrate analogue for phytate whichwould offer the possibility of rapid screening pro-cedures (Berry and Berry, 2005).

An interesting ecosystem that needs to bemore thoroughly investigated is the gastrointesti-nal tract of animals. Yanke et al. (1998, 1999)reported the isolation of anaerobic microorgan-isms (e.g. Selenomona ruminatium) from the rumenfluid of cows and suggested their role in phytateutilization. There is evidence that metal complex-ation in cattle fed a grain-based diet can result inmanures that contain appreciable concentrationsof inositol phosphates (see Dao, Chapter 11, thisvolume). However, it is generally considered thatcattle do not excrete significant amounts of inosi-tol phosphates (see Leytem and Maguire,Chapter 10, this volume). Possible explanationsare that ruminants contain a pH-neutral foregutthat is effective for phytate degradation, or thathydrolysis of inositol phosphates occurs underanaerobic–fermentative conditions. Plant-derivedphytases in feed may also be active in the rumen(see Lei and Porres, Chapter 9, this volume).Monogastric animals, on the other hand, havefermentative digestion after an acidic stomach,which may either be detrimental to incomingphytate-degrading bacteria or denature phytases.Thus, poultry manures tend to contain largeamounts of phytate (e.g. McGrath et al., 2005). Inspite of this, there is some evidence indicating thatbacteria in the hindgut of monogastric animalsmight also hydrolyse phytate, resulting in adecrease in the excreta of swine and poultry(Leytem et al., 2004). Further work to understandthe degradation of phytate in the gut of bothmonogastric animals and ruminants is required.

Ecological understanding of phytate-degrading microorganisms

In understanding the potential for degradation ofinositol phosphates in terrestrial and aquaticsystems, knowledge of the geochemistry of the envi-ronment is required. Microorganisms live in niche


environments, so any gross parameters measuredshould be considered in this context. Interactionbetween microorganisms and inositol phosphatesrequires that the substrate be available to themicroorganism, although this can be influenced bya range of physiological and abiotic processes.Factors affecting the solubility of inositol phos-phates in soil include pH, clay content, and thepresence and concentration of metal ions in soilsolution (reviewed by Turner et al., 2002; seeTurner, Chapter 12, and Celi and Barberis,Chapter 13, this volume). In particular, complexa-tion of inositol phosphates with metals such as ironand aluminium can render the resulting insolublephytates unavailable to microorganisms (e.g.Greenwood and Lewis, 1977). Microorganismsalso require carbon and nitrogen and suitablephysiological conditions such as pH and moisture,although phosphorus rarely limits microbial growth( Jakobsen et al., 2005). An exception may be freshwater planktonic cells (Schindler, 1971; O’Sullivan,1992; McComb and Davis, 1993), but in most soil-based environments the concentration of phos-phate, or organic phosphates other than inositolphosphates, may provide sufficient phosphorus tosupport microbial growth.

Molecular approaches to the ecology ofphytate-degrading microorganisms

Culture-independent approaches based on DNAtechnology indicate that generally <1% of microor-ganisms, particularly bacteria, can be cultured fromenvironmental samples (Kaeberlein et al., 2002). Ofthose that have been cultured and characterized,few have been assessed for their ability to degradephytate. It is therefore unclear whether the abilityto utilize phytate is a common trait or restricted toselect genera (e.g. as listed in Table 5.3). However,given the level of diversity present in phytate-degrading microorganisms so far isolated, and thevast number of microorganisms in environmentalsamples that have yet to be cultured, it seems likelythat the number of microorganisms able to degradephytate is large. Development of specific moleculartools to analyse environmental samples could beused to address this issue.

Molecular community diversity analysis ofecosystems can be conducted using DNA ampli-fication techniques based on the polymerasechain reaction (PCR). These are particularlyuseful when applied to the 16S ribosomal gene,

which is ubiquitous and has highly conservedsequence domains across the eubacterial andarchaebacterial kingdoms. Techniques such asterminal-restriction fragment length polymor-phism (T-RFLP) and denaturing gradient gelelectrophoresis (DGGE) can be used to investi-gate the structure of microbial communities inenvironmental samples and identify ‘shifts’ inmajor groups of bacteria (Tajima et al., 2001;Possemiers et al., 2004; Van der Gucht et al.,2005). Such approaches could be applied toenvironment samples where concentrations ofinositol phosphates are known to change (e.g.after the addition of manures to soil) to identifykey groups of microorganisms that might beinvolved in their hydrolysis. A key challenge forfuture studies is to link changes in microbialcommunity structure to specific functionalgroups of bacteria and/or specific biochemicalprocesses within the environment.

Molecular-based techniques can also beused to identify the presence (and ideally thefunction) of specific genes. To study gene shifts,sufficient DNA homology among target genes isrequired and at present this may be possible withthe β-propeller phytases. This family of phytaseshas been shown (Cheng and Lim, 2006) to occurin a wide range of bacteria isolated from diverseenvironments (e.g. as identified using online dataof the National Center for Biotechnology Infor-mation, available at www.ncbi.nlm.nih.gov).Preliminary work has shown that Shewanella onei-densis MR-1 phytase shares amino acid homologywith a Shewanella organism in sea water samplesfrom the Sargasso Sea (Venter et al., 2004). Basedon this assessment and enzyme kinetics of theS. oneidensis phytase, it is possible that Shewanellaplays a significant role in the degradation of phy-tate that has been observed in marine environ-ments (Suzumura and Kamatani, 1995b; Chengand Lim, 2006). An alternative approach is to usesets of β-propeller-specific nucleic acid primers toprobe microbial communities and identify, andpossibly isolate, microorganisms with homolo-gous genes ( J. Hill, 2006, unpublished data).

Conclusions and Future Research

It is becoming clear that microorganisms withthe ability to utilize inositol phosphates are ubiqui-tous in both terrestrial and aquatic environments.


www.ncbi.nlm.nih.gov

This challenges the conventional perception thatinositol phosphates are recalcitrant and do notcontribute greatly to the nutrition of organisms.The ecological implications of this abundance ofphytate-degrading microorganisms remainlargely unexplored, but are likely to be significantin several environments. For example, inositolphosphates can no longer be considered unim-portant in terms of water quality (see McKelvie,Chapter 16, this volume).

Assessment of phytate-degrading microor-ganisms requires screening (usually starting withsolid or liquid enrichment), identification of theorganism responsible, purification of the enzyme,and evaluation of growth parameters and enzymeproperties. While current protocols are labour-intensive, they provide important informationconcerning the potential of different groups ofmicroorganisms to degrade phytate and, increas-ingly, of their ecological significance. However,much remains to be achieved. The use of higherthroughput systems (e.g. robotic sampling and

culturing systems) should allow a wider diversityof environments to be investigated in detail.

Irrespective of this, it is important that effortsto analyse microbial populations for phytatedegradation also consider the environment inwhich the microorganisms operate. While thedevelopment of new screens that adopt techniquessuch as community and gene analysis will clearlyassist in this process, these approaches must adoptmeaningful ways to link the analysis of commu-nity structure with microbial function and ecosys-tem characteristics. Molecular approaches willneed to incorporate analytical tools such as 31PNMR spectroscopy to determine the speciationand concentration of inositol phosphates. Unitingmolecular and analytical tools is complex and willrequire collaborative expertise. However, until thedevelopment of more gene- and enzyme-specificapproaches, these are important ways to gain anecological understanding of phytate-degradingorganisms and their role in the utilization of inos-itol phosphates in the environment.


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Wyss, M., Pasamontes, L., Friedlein, A., Remy, R., Tessier, M., Kronenberger, A., Middendorf, A., Lehmann, M.,Schnoebelen, L., Rothlisberger, U., Kusznir, E., Wahl, G., Muller, F., Lahm, H.W., Vogel, K. and van Loon,A. (1999b) Biophysical characterization of fungal phytases (myo-inositol hexakisphosphate phosphohydrolases):molecular size, glycosylation pattern, and engineering of proteolytic resistance. Applied and EnvironmentalMicrobiology 65, 359–366.

Xiong, A.S., Yao, Q.H., Peng, R.H., Li, M., Fan, H.Q., Guo, M. J. and Zhang, S. (2004) Isolation, characteriza-tion, and molecular cloning of the cDNA encoding a novel phytase from Aspergillus niger 113 and high expres-sion in Pichia pastoris. Journal of Biochemistry and Molecular Biology 37, 282–291.

Yamada, K., Minoda, Y. and Yamamoto, S. (1968) Phytase from Aspergillus terreus. I. Production purification andsome general properties of enzyme. Agricultural and Biological Chemistry 32, 1275–1282.

Yanke, L. J., Bae, H.D., Selinger, L.B. and Cheng, K. J. (1998) Phytase activity of anaerobic ruminal bacteria.Microbiology 144, 1565–1573.

Yanke, L. J., Selinger, L.B. and Cheng, K. J. (1999) Phytase activity of Selenomonas ruminantium: a preliminary char-acterization. Letters in Applied Microbiology 29, 20–25.

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©CAB International 2007. Inositol Phosphates: Linking Agriculture and the Environment78 (eds B.L. Turner, A.E. Richardson and E.J. Mullaney)

6 Phytate-degrading Enzymes:Regulation of Synthesis inMicroorganisms and Plants

Ralf GreinerFederal Research Centre for Nutrition and Food, Centre for Molecular Biology,

Haid-und-Neu-Strabe 9, D 76131 Karlsruhe, Germany

Phytate-degrading enzymes, also known asphytases, have a wide distribution in plants,microorganisms and some animal tissues(Konietzny and Greiner, 2002; see Hill andRichardson, Chapter 5, and Mullaney andUllah, Chapter 7, this volume). They belong toa special class of phosphomonoesterases termedmyo-inositol hexakisphosphate phosphohy-dolases, which are capable of initiating the step-wise release of phosphate residues from phytate(salts of myo-inositol hexakisphosphate), themajor storage form of phosphate in plant seedsand pollen. The ability of such enzymes tohydrolyse phytate is usually known only fromin vitro assays, and information on their in vivofunction is rather limited. As enzymes are classi-fied in general by their in vivo function, the term‘phytate-degrading enzyme’ is preferred here tothe term ‘phytase’, and is used throughout thechapter. The classification of enzymes as phy-tases based solely on in vitro assays becomesproblematic when considering enzymes such asglucose-1-phosphatase in Escherichia coli andEnterobacter cloacae. This enzyme can hydrolysephytate, albeit slowly, in vitro even though this isclearly not its in vivo function. Unless otherwisestated, designation of an enzyme as ‘phytate-degrading’ is based on in vitro assay using solublesodium phytate.

Phytate-degrading Enzymes andTheir Classification

Phosphomonoesterases are a diverse group ofenzymes that encompass a range of sizes, struc-tures and catalytic mechanisms. Based on theamino acid residue in the active site, phytate-degrading enzymes can be referred to as histidineacid phosphatases, β-propeller phosphatases, cyst-eine phosphatases and purple acid phosphatases(see Mullaney and Ullah, Chapter 7, this volume).Two classes of phytate-degrading enzymes arerecognized by the International Union of Pureand Applied Chemistry and the InternationalUnion of Biochemistry (IUPAC–IUB): 3-phytase(EC 3.1.3.8), which initially removes phosphatefrom the D-3 position of the myo-inositol ring, and6-phytase (EC 3.1.3.26), which preferentially initi-ates phytate dephosphorylation at the L-6 (D-4)position. However, phytate-degrading enzymesinitiating phytate degradation at the D-5 and D-6positions, respectively, have been found in nature(Barrientos et al., 1994; Greiner et al., 2000a).Phytate-degrading enzymes from microorganismsare considered to be 3-phytases, whereas 6-phytases are said to be characteristic of theseeds of higher plants.

Most of the phytate-degrading enzymesstudies so far with respect to their pathway of

phytate degradation fit into this consideration(Fig. 6.1). However, this is not a general rule, asexemplified by the indication of 3-phytase activ-ity in lupine (Greiner et al., 2002) and soybeanseeds (R. Greiner, 2000 unpublished data) and 6-phytase activity in Paramecium (van der Kaay andvan Haastert, 1995) as well as E. coli (Greineret al., 2000a). However, it is worth mentioningthat the 6-phytase activity of plant seeds initiallyhydrolyses the L-6 (D-4) phosphate residue fromphytate, whereas phytate-degrading enzymesfrom Paramecium and E. coli initially remove thephosphate residue attached to the D-6 (L-4) pos-ition. Phytate-degrading enzymes are found inmultiple forms, especially in plant seeds. Theseforms may even exhibit different stereospecificity ofmyo-inositol hexakisphosphate dephosphorylation,as reported recently for the phytate-degradingenzymes from lupines (Greiner et al., 2002).

The phosphate residues of phytate arereleased by phytate-degrading enzymes at differentrates and in different order. Independent of theirbacterial, fungal or plant origin, the majority ofthe phytate-degrading enzymes exhibiting an opti-mum for phytate hydrolysis under acidic pH con-ditions release five of the six phosphate residues ofphytate, and the final degradation product wasidentified as myo-inositol 2-phosphate (Konietznyand Greiner, 2002). Dephosphorylation of myo-inositol 2-phosphate occurs only in the presence ofhigh enzyme concentration during prolongedincubation. After removal of the first phosphateresidue from phytate these histidine acid phytate-degrading enzymes continue dephosphorylationadjacent to a free hydroxyl group. The majorphytate degradation pathways of 6-phytases ofplant origin and 3-phytases differ only in the myo-inositol pentakisphosphate intermediate generated(Fig. 6.1). The two exceptions reported so far arethe 6-phytases from mung bean (Maiti et al., 1974)and wheat F2 phytase (Lim and Tate, 1973). Themicrobial 6-phytases, however, generate a com-pletely different set of myo-inositol phosphate inter-mediates (Fig. 6.1). In addition, an acidphosphatase with phytate-degrading activity wasidentified in members of the Enterobacteriaceae fam-ily, such as E. coli (Cottrill et al., 2002), Pantoeaagglomerans (Greiner, 2004) and E. cloacae(R. Greiner, 2005 unpublished data), whichpreferably degrades glucose1-phosphate. Theseenzymes were shown to hydrolyse only the phos-phate residue at the D-3 position of phytate, pro-

ducing D-myo-inositol 1,2,4,5,6-pentakisphosphateas the sole hydrolysis product.

The alkaline phytate-degrading enzymes fromcattail (Typha spp.; Hara et al., 1985), lily pollen(Lilium longiflorum; Barrientos et al., 1994) andBacillus subtilis (Kerovuo et al., 2000) yield myo-inosi-tol trisphosphate as the final product of phytatedephosphorylation (Fig. 6.2). With the exception ofthe phytate-degrading enzyme from L. longiflorum,alkaline phytate-degrading enzymes represent theclass of β-propeller phosphatases. These seem toprefer the hydrolysis of every second phosphaterather than of adjacent ones, generating myo-inosi-tol 2,4,6-trisphosphate and myo-inositol 1,3,5-trisphosphate as the final dephosphorylationproducts (Kerovuo et al., 2000). Recent studies onthe phytate-degrading enzymes of B. subtilis 168, B.amyloliquefaciens ATCC 15841 and B. amyloliquefa-ciens 45, however, point to myo-inositol 2,4,6-trisphosphate as the sole final product of phytatedegradation (R. Greiner, 2005 unpublished data).The alkaline phytate-degrading enzyme from L.longiflorum possesses the conserved active site motifscharacteristic of histidine acid phosphatases (Mehtaand Murthy, 2005) and generates a myo-inositoltrisphosphate as the final phytate dephosphoryla-tion product in a single degradation pathway bypreferring removal of adjacent phosphate groups(Barrientos et al., 1994).

Regulation of phytate-degradingenzyme formation in microorganisms

and plants

In plant seeds and microorganisms, constitutiveas well as inducible phytate-degrading enzymeshave been identified. In microorganisms, expres-sion of the inducible phytate-degrading enzymesis subjected to a complex regulation, but theirformation is not controlled uniformly among dif-ferent microorganisms (Konietzny and Greiner,2004). Until now, phytate-degrading enzymeproduction was studied in some detail only inE. coli (Greiner et al., 1993), Raoultella terrigena(Greiner et al., 1997; Zamudio et al., 2002) andSaccharomyces cerevisiae (Andlid et al., 2004). A largeincrease in phytate-degrading activity wasreported in germinating plant seeds and pollen,but the biochemical mechanisms leading to thisrise in enzyme activity is still not well understood.

Synthesis in Microorganisms and Plants 79

80 R. Greiner

I(1,2,3,4,5,6)P6

D-I(1,2,3,5,6)P5 D-I(1,2,4,5,6)P5 D-I(1,2,3,4,5)P5

D-I(1,2,3,6)P4 D-I(1,2,5,6)P4 D-I(2,3,4,5)P4 D-I(1,2,3,4)P4

I(1,2,3)P3 D-I(1,2,6)P3 D-I(2,4,5)P3 I(1,2,3)P3

D-I(1,2)P2 I(2,5)P2 D-I(2,3)P2

I(2)P

ParameciumE. coli appA

Barley P1, barleyP2, spelt D12,wheat PHY1,wheat PHY2,oat, rice, rye,faba bean, lupineLP2

Wheat F2,mung bean(1)

E. coli appA,Paramecium

Pseudomonas, Saccharomycescerevisiae, Aspergillus ficuum,lupine LP11, lupine LP12,Pantoea agglomerans (2),Escherichia coli agp (2),Eenterobacter cloacae (2)

Barley P1, barley P2, speltD12, wheat PHY1, wheatPHY2, wheat F2, oat, rice,rye, faba bean, lupineLP2, mung bean

Fig. 6.1. Major phytate degradation pathways by acid phytate-degrading enzymes barley P1, barley P2,spelt D12, rye, oat (Greiner and Larsson Alminger, 2001); wheat PHY1, wheat PHY2 (Nakano et al.,2000); rice (Hayakawa et al., 1990); faba bean, lupine LP11, lupine LP12, lupine LP2 (Greiner et al.,2002); wheat F2 (Lim and Tate, 1973); mung bean (Maiti et al., 1974); Saccharomyces cerevisiae(Greiner et al., 2001); Pseudomonas (Cosgrove, 1970); Escherichia coli (Greiner et al., 2000a; Cottrillet al., 2002); Paramecium (van der Kaay and van Haastert, 1995); Aspergillus ficuum (Chen and Li,2003); Pantoea agglomerans (Greiner, 2004); Enterobacter cloacae (R. Greiner, 2005 unpublished data).

(1) Generates also D-I(1,2,6)P3 and D-I(1,2)P2 as intermediates(2) D-I(1,2,4,5,6)P5 is the final product of phytate dephosphorylation

Regulation of phytate-degrading enzymeformation in microorganisms

In non-limiting media, formation of the majorityof the bacterial phytate-degrading enzymes wasturned off in exponentially growing cells andstarted as soon as the cultures entered the station-ary phase (Shimizu, 1992; Greiner et al., 1993,1997; Sreeramulu et al., 1996; Choi et al., 1999;Zamudio et al., 2001; De Angelis et al., 2003). Inmoulds, phytate-degrading enzyme formationwas growth-associated (Vats and Banerjee, 2002).Enzyme activity started to increase from thebeginning of growth and continued to increaseup to the onset of the stationary phase.

As synthesis of bacterial phytate-degradingenzymes started as soon as the growth rate beganto fall, it was suggested that either a nutrient or anenergy limitation, both known to occur in the sta-tionary phase, could cause their induction. Amongthe nutrient limitations tested, only carbon starva-tion was able to provoke an immediate synthesis ofa phytate-degrading enzyme in R. terrigena (Greiner

et al., 1997). However, phytate-degrading enzymeformation in E. coli was triggered by phosphatestarvation, while carbon, nitrogen and sulphur lim-itation were ineffective (Touati et al., 1987). A tightregulatory inhibition of the formation of phytate-degrading enzymes by phosphate levels was gener-ally observed in all microorganisms, includingmoulds, yeast and bacteria, with the exception ofR. terrigena (Greiner et al., 1997) and the rumen bac-teria (Yanke et al., 1998), but only rarely couldmicroorganisms utilize phytate as the sole source ofcarbon and phosphate. However, a small amountof phosphate in the growth medium probably stim-ulates phytate-degrading enzyme formation byenhancing microbial growth. Phosphate was shownto exert its effect on the synthesis of phytate-degrading enzymes at the level of transcription.The repression by phosphate seems to be less sig-nificant with more complex media (Greiner andFarouk, 2005), although it is not known which spe-cific media components could account for this.

Expression of phytate-degrading enzymesalso depends on the nature of the carbon source,


I(1,2,3,4,5,6)P6

D-I(1,2,3,4,6)P5 D/L-I(1,2,3,4,5)P5 D/L-I(1,2,4,5,6)P5 D-I(1,2,4,5,6)P5

D/L-I(1,2,3,4)P4 I(1,2,3,5)P4 I(2,4,5,6)P4

I(1,2,3)P3 I(1,3,5)P3 I(2,4,6)P3

B. subtilis 168,B. amyloliquefaciensATCC 15841,B. amyloliquefaciens 45

Bacillussubtilis

Lily

Fig. 6.2. Major phytate degradation pathways by the alkaline phytate-degrading enzymes Bacillus subtilis(Kerovuo et al., 2000); B. subtilis 168, B. amyloliquefaciens ATCC 15841, B. amyloliquefaciens 45 (fromR. Greiner, 2005 unpublished data); lily (from Barrientos et al., 1994). The final phytate degradationproduct of cattail phytase was identified as myo-inositol trisphosphate (Hara et al., 1985), but noinformation was provided about the configuration of the generated phytate degradation products.

the initial pH and the temperature used for culti-vation of the microorganisms. Both biomass andenzyme production respond to all these param-eters, but temperature and pH for maximalgrowth and production of phytate-degradingenzymes were shown to be different in somemoulds and yeast (Lambrechts et al., 1993; Kimet al., 1999; Sano et al., 1999; Mandviwala andKhire, 2000; Andlid et al., 2004), indicating a bio-mass-independent effect of temperature and pHon the synthesis of phytate-degrading enzymes inthese microorganisms. No comparable studieshave been performed so far in bacteria. In thepresence of simple, easily fermentable sugars,strong repression of the formation of phytate-degrading enzymes was observed in severalmicroorganisms, including moulds, yeast and bac-teria (Shieh and Ware, 1968; Han and Gallagher,1987; Lambrechts et al., 1993; Sano et al., 1999;Lan et al., 2002; Vats and Banerjee, 2002; Vohraand Satyanarayana, 2002).

As with phosphate, an optimal concentrationof glucose is also required. Low glucose levels resultin low phytate-degrading activity due to reducedbiomass production, whereas high levels inhibitenzyme production. However, the presence of glu-cose causes high levels of phytate-degrading activityin E. coli (Touati et al., 1987) and Lactobacillusamylovorus (Sreeramulu et al., 1996). The regulationof the formation of phytate-degrading enzymes wasnot suggested to be due to the carbon source itself,but to a change in the level of cellular cyclic adeno-sine monophosphate (cAMP). It has been estab-lished that a complex of cAMP with a proteincalled cAMP receptor protein (CRP) plays a cen-tral role in activating and repressing the expressionof many genes (Saier et al., 1996). Cellular cAMPlevels depend upon cell physiology, including thecarbon source in the growth medium. ThatcAMP–CRP is directly involved in the regulationof the formation of phytate-degrading enzymes wasshown in E. coli (Touati et al., 1987) and R. terrigena(Zamudio et al., 2002). In R. terrigena, synthesis ofthe phytate-degrading enzyme was downregulatedby cAMP–CRP in the stationary phase and upreg-ulated during exponential growth. Synthesis ofphytate-degrading enzymes in E. coli has beenreported to be downregulated by cAMP–CRPunder all growth conditions. In moulds such asAspergillus niger, formation of mycelial pellets in thepresence of glucose or fructose as the sole carbonsource was shown to be responsible for the low

enzyme yields (Shieh and Ware, 1968; Han andGallagher, 1987). Dispersed growth and thereforean increase in phytate-degrading enzyme produc-tion could be obtained by using a medium contain-ing a surfactant.

For several Klebsiella spp. it was reported thatphytate is needed to induce phytate-degradingenzyme production (Shah and Parekh, 1990;Tambe et al., 1994; Greiner et al., 1997), but thebacteria were unable to grow on phytate as thesole carbon source. Substrate induction was alsofound for Mitsuokella jalaludinii (Lan et al., 2002)and several Bacillus spp. (Powar and Jagannathan,1982; Kerovuo et al., 1998; Kim et al., 1998),whereas phytate had no effect on the formation ofphytate-degrading enzymes in E. coli (Greineret al., 1993), Arxula adeninivorans (Sano et al., 1999)and Selenomonas ruminantium (Yanke et al., 1998).A reduced synthesis of the phytate-degradingenzymes in the presence of phytate was observedeven in Schwanniomyces castellii (Lambrechts et al.,1993). An increased phosphate concentration inthe growth medium due to phytate hydrolysis bythe phytate-degrading enzymes secreted by theyeast could be responsible for this phenomenon.This suggestion would be in agreement with theobservation that wheat and rice bran are excellentsubstrates for the production of extracellular phy-tate-degrading enzymes in microorganisms. Asphytate in bran is less soluble than sodium phy-tate, phosphate concentrations are lower due to aslower release from bran phytate, and thereforerepression of enzyme synthesis by phosphate isreduced. This ensures a continuous production ofphytate-degrading enzymes during the whole fer-mentation process.

The formation of phytate-degrading enzymesin Pseudomonas spp. (Irving and Cosgrove, 1971)and Klebsiella aerogenes (Tambe et al., 1994) wasreported to be significantly induced in the pres-ence of myo-inositol as the sole carbon source,although this was less effective than phytate. In theother Klebsiella spp. studied myo-inositol was ineffec-tive (Shah and Parekh, 1990; Greiner et al., 1997).Anaerobiosis was effective in inducing phytate-degrading enzyme formation in E. coli (Greineret al., 1993) and S. castellii (Lambrechts et al., 1993),whereas aeration had a positive effect on the pro-duction of phytate-degrading enzymes in A. ficcum(Nair et al., 1991).

The presence of calcium ions in the growthmedium was found to result in higher extracellular

82 R. Greiner

phytate-degrading activity in Bacillus spp.(Shimizu, 1992; Choi et al., 1999). However, themetal ions are not supposed to induce enzymeexpression, but to stabilize the secreted enzyme.Binding of two calcium ions to high-affinity cal-cium binding sites was shown to result in adramatic increase in thermostability of the phy-tate-degrading enzymes from Bacillus by joiningloop segments remote in the amino acid sequence.Binding of three additional calcium ions to low-affinity calcium binding sites at the top of theenzyme molecule turns on the catalytic activity ofthe enzyme by converting the highly negativelycharged cleft into a favourable environment forthe binding of phytate (Shin et al., 2001).Regulation of phytate-degrading enzyme forma-tion on a molecular level was studied in detail inE. coli and S. cerevisae only.

Regulation of phytate-degrading enzymeformation in Escherichia coli

At least two different phosphatases located in theperiplasma of E. coli are capable of accepting myo-inositol hexakisphosphate as a substrate. The agp-encoded acid phosphatase (EC 3.1.3.10)hydrolyses only the D-3 phosphate residue fromphytate to produce D-myo-inositol 1,2,4,5,6-pen-takisphosphate as the sole hydrolysis product(Cottrill et al., 2002). This enzyme has broad sub-strate specificity for phosphorylated compoundsbut demonstrates its highest activity towards glu-cose 1-phosphate, and it is believed that its func-tion is to scavenge glucose (Pradel and Boquet,1991). It appears to be largely synthesized consti-tutively, although small effects have been notedin regard to the amounts of this enzyme madeunder various growth conditions (Wanner, 1996).

The appA-encoded phosphatase (EC3.1.3.26) is highly specific for phytate andsequentially removes five of the six phosphategroups, starting with that attached to the D-6position of the myo-inositol ring (Greiner et al.,2000a). Accumulation of this E. coli enzyme iscontrolled at the level of transcription and itsexpression is regulated by a complex regulatorymechanism involving several factors (Fig. 6.3).The appCBA operon contains the genes appA aswell as cbdA and cbdB, coding for a putativecytochrome oxidase. These three genes are co-

transcribed from a promoter (appC) locatedimmediately upstream of cbdA and the operon, inaddition, contains an internal promoter (appA)positioned upstream of appA (Atlung andBrøndsted, 1994). Both promoters are induced byphosphate starvation and by entry into the sta-tionary phase (Atlung and Brøndsted, 1994), butthe induction of the appC promoter is muchstronger than that of the appA promoter. Theinduction by phosphate starvation has beenshown to be independent of the PHO regulon(Touati et al., 1987), which is involved in the scav-enging and specific uptake of phosphate fromextracellular sources. No information on the reg-ulation of the appA promoter under phosphatestarvation and upon entry into stationary phase isavailable. In addition, the appC promoter wasfound to be activated by anaerobic growth condi-tions (Atlung and Brøndsted, 1994) and carbonstarvation (Atlung et al., 1997). At low glucoselevels the cAMP–CRP complex was suggested todirectly interact with the appA promoter (Touatiet al., 1987).

Transcription from the appC promoter isdependent on the σS subunit of the RNA poly-merase under all growth conditions tested, specif-ically during exponential growth, entry into thestationary phase in rich medium, starvation forcarbon and phosphate and upon osmotic upshift(Atlung et al., 1997). It was suggested that σS

affects expression of the appCBA operon directly.That σS controls the expression of genes respond-ing to starvation, and cellular stress is well estab-lished. Therefore, the intracellular concentrationof σS present in E. coli is influenced strongly byenvironmental factors. It increases upon entryinto the stationary phase in rich medium, duringstarvation for carbon, nitrogen and phosphate(Gentry et al., 1993; Lange and Hengge-Aronis,1994), and is increased strongly by osmoticupshift (Muffler et al., 1996). Although σS alwaysaffects the level of expression from the appC pro-moter, it is considered to have a regulatory roleonly during induction by osmotic upshift andupon entry into the stationary phase in richmedium, but the phosphate starvation-inducedincrease in σS concentration is not involved inthe regulation of this operon.

The expression of σS-dependent genes doesnot depend solely on the concentration of σS inthe cell, but additional regulatory factors differ-entially modulate the expression of these genes.


Many σS-dependent genes are also under positiveor negative control of other well-known globalregulators such as cAMP–CRP, ArcA, AppY andFnr. The appCBA operon was shown to be a tar-get for the transcriptional activator AppY(Brøndsted and Atlung, 1996). The induction ofthe appC promoter by anaerobiosis is fullydependent on AppY, whereas its induction byphosphate starvation and upon entry into station-ary phase (Fig. 6.3) is dependent strongly, but notexclusively, on this transcriptional regulator.AppY has no effect on the induction of the appApromoter. AppY expression itself is induced byanaerobiosis, starvation for phosphate and car-bon, and upon entry into the stationary phase(Fig. 6.3), indicating that the increased AppYconcentration under these conditions contributesto the increased expression of this E. coli phos-phatase. An increase in σS was shown to beinstrumental in the induction of appY during car-bon starvation (Fig. 6.3).

The stationary phase induction of appY isonly partially dependent on σS, whereas theinduction during phosphate starvation and thegrowth rate regulation is independent of σS

(Fig. 6.3). AppY-dependent induction of theappCBA operon during anaerobiosis and phos-phate starvation seems to be mediated by a com-bination of increased appY expression and anactivating signal for AppY generated under bothconditions (Fig. 6.3). Formate, an intermediate inthe mixed acid fermentation, was suggested to bethis activating signal (Brøndsted and Atlung,1996). However, the appC promoter also seems tobe stimulated quite efficiently by AppY undernon-activating conditions. In addition, appYexpression always increased when glycerol wasused as the carbon source. This was not due to apositive regulation by cAMP–CRP, but due to anincrease in the doubling time in the presence ofglycerol compared to glucose as a carbon source.That expression of appY is inversely correlated

84 R. Greiner

cBcB

Cell membrane

PappC PappAcbdA cbdB appA

AppY

AppY

XPappY appY

Activa

tion

Weakinduction

Stronginduction

Phytase

mRNA

mRNA

ArcA

P

ArcB

Activation byphosphorylation

Induction

Fig. 6.3. Regulation of phytate-degrading enzyme formation in Escherichia coli. PappC = appC promoter;PappA = appA promoter; PappY = appY promoter; cbdA, cbdB = genes coding for a putative cytochromeoxidase; appA = gene encoding a phytate-degrading enzyme; appY = gene encoding AppY; AppY,ArcA = transcriptional regulators; ArcB = oxygen sensor; X = formate; P = phosphate.

with the growth rate was shown previously(Brøndsted and Atlung, 1996).

The ArcA response regulator, a second tran-scriptional regulator that is activated by the ArcBsensor in response to reduced respiration, acti-vates transcription of the appCBA operon duringentry into stationary phase and under anaerobicgrowth conditions (Fig. 6.3; Brøndsted andAtlung, 1996). During stationary phase induc-tion, much of the ArcA effect is by AppY. It ispossible that only the weak and early induction ofthe appCBA operon is mediated by ArcA in anAppY-independent manner. The signal that leadsto the induction of the appC promoter upon entryinto stationary phase may be primarily due tooxygen deprivation caused by an increase in celldensity. AppY and ArcA depend on each otherwhen activating the transcription of the appCBAoperon during anaerobic growth. AppY expres-sion was induced immediately by anaerobiosis, aprocess that is dependent on ArcA. The expres-sion of the appCBA operon does not respondimmediately to anaerobiosis but is delayed onegeneration, possibly due to the lack of sufficientAppY at the onset of anaerobiosis.

Electron acceptors, which can be used inanaerobic respiration, repress the expression ofthe appCBA operon (Brøndsted and Atlung, 1996).The repression is particularly pronounced in thepresence of nitrate. Since appY expression was notaffected significantly by anaerobic energy metabo-lism (i.e. fermentation vs. anaerobic respiration),induction of the appCBA operon cannot be medi-ated by changes in the expression of AppY. It wassuggested that nitrate repression was partiallydependent on NarL, which activates transcriptionof operons involved in nitrate respiration andrepresses the synthesis of alternate respiratoryenzymes (Berg and Stewart, 1990). The residualnitrate repression could be mediated by NarP(Rabin and Stewart, 1993). Alternatively, theresidual anaerobic repression by nitrate and therepression by fumarate could be indirect effects ofArcA, as the level of active ArcA is dependent onthe respiratory state of the cell.

Regulation of phytate-degrading enzymeformation in Saccharomyces cerevisiae

In the yeast S. cerevisiae, the PHO regulon controlsexpression of the PHO genes at the transcription

level depending on the extracellular phosphateconcentration (Lemire et al., 1985), but it is notknown how extracellular phosphate levels aredetected by the yeast. The PHO gene family isinvolved in the scavenging and specific uptake ofphosphate from extracellular sources and at leastthree phosphatases, encoded by PHO5, PHO10and PHO11, are capable of hydrolysing myo-inos-itol hexakisphosphate (Andlid et al., 2004). Allthese enzymes are extracellular oligomeric glyco-proteins with an acidic pH optimum and broadsubstrate specificity. PHO5 encodes the majorcontributor to the secreted acid phosphataseactivity, whereas PHO10 and PHO11 encode onlya minor fraction (Lemire et al., 1985).

The information about the extracellularphosphate level is transmitted to the phosphataseencoding genes by a set of positive and negativeregulatory proteins, which are encoded by at leastthe following genes: PHO2, PHO4, PHO80,PHO81 and PHO85. Besides PHO5, PHO10 andPHO11, at least the following additional genes areknown to be regulated by the PHO regulon: PHO8encoding a non-specific repressible alkaline phos-phatase localized in the vacuole; PHO84 andPHO89 encoding high-affinity phosphate trans-porters localized in the plasma membrane; GIT1encoding a glycerophosphoinositol transporterlocalized in the plasma membrane; PHO86 encod-ing a protein required for the correct localizationof Pho84p in the plasma membrane; PHO81,SPL2 and YPL110C encoding regulatory proteins,and several genes encoding various phosphatemetabolism enzymes such as the PHM genes thatare involved in polyphosphate synthesis or degra-dation (Pinson et al., 2004).

For the transcriptional activation of PHO5,PHO10 and PHO11 the two DNA-binding pro-teins Pho4p, encoded by PHO4, and Pho2p,encoded by PHO2, are needed (Fig. 6.4). Pho4pbinds to a specific cis-acting regulatory site(UAS) in the promoter of all PHO genes(Oshima, 1997), and Pho2p forms a ternarycomplex with Pho4p on a PHO promoter(Barbaric et al., 1996). However, Pho2p does nothave a direct function in the transduction of theextracellular phosphate levels, but interaction ofthe Pho2p/Pho4p/UAS ternary complex withbasal transcription factors is considered to initi-ate transcription of the PHO genes (Magbanuaet al., 1997). Both PHO4 and PHO2 are tran-scribed at low levels (Yoshida et al., 1989).


PHO4 transcription is constitutive, whereasPHO2 transcription is self-regulated.

High extracellular phosphate concentrationsresult in phosphorylation of five serine residues inPho4p by a complex of the negative regulatorsPho80p and Pho85p (Kaffman et al., 1994). Thisphosphorylated Pho4p is not imported into thenucleus (O’Neill et al., 1996) and therefore tran-scription of PHO genes is turned off (Fig. 6.4).Pho81p, which is inactive under high extracellu-lar phosphate concentrations, is activated whenthe extracellular phosphate level is sufficientlylow. Activated Pho81p inhibits the function ofthe Pho80p–Pho85p complex (Ogawa et al.,1995), thus allowing translocation of Pho4p intothe nucleus, where Pho4p, together with Pho2p,activates the transcription of the PHO genes(Fig. 6.4; Komeili and O’Shea, 1999). PHO80and PHO85 are transcribed constitutively at lowlevels (Madden et al., 1990), whereas transcription

of PHO81 is regulated by the level of extracellularphosphate by the same PHO regulon, indicatingthat the regulatory circuit forms a positive feed-back loop (Ogawa et al., 1993).

Regulation of phytate-degradingenzyme formation in plants

Studies in plant seeds and pollen indicate thatthere are constitutive and germination-induciblephytate-degrading enzymes. Thus, two mainmechanisms appear to be involved in the regula-tion of phytate breakdown during germination:control of the activity of the hydrolytic enzymesand control of their rate of synthesis. Constitutivephytate-degrading enzymes, which are found inall seeds and pollen, are present in a fully activeform in mature seeds and pollen, and are consid-

86 R. Greiner

PPHO

Nuclearmembrane

Pho4p

Cell membranePho89p

Pho85pkinase

Pho80pcyclase

Pho81pinhibitor

mRNAPho4p

Phytase

Pho2p

PHO-regulated gene

Pho2p

Pho84p

Pho89p

Pho84p

Fig. 6.4. Regulation of phytate-degrading enzyme formation in Saccharomyces. PPHO = PHO promoter;Pho2p, Pho4p = regulatory proteins (DNA-binding proteins); Pho81p = regulatory protein (inhibitor);Pho80p = cyclase; Pho85p = kinase; Pho84p, Pho89p = high-affinity phosphate transporters; PHO -regulated genes = PHO5, PHO10 and PHO11 encoding repressible non-specific acid phosphatases;PHO8 encoding a non-specific repressible alkaline phosphatase; PHO84 and PHO89 encoding high-affinity phosphate transporters; GIT1 encoding a glycerophosphoinositol transporter; PHO86 encoding aprotein required for the correct localization of Pho84p; PHO81, SPL2 and YPL110C encoding regulatoryproteins; several genes encoding various phosphate metabolism enzymes such as the PHM genes.

ered to start phytate breakdown during imbibi-tion. Germination-inducible phytate-degradingenzymes are synthesized de novo either from along-lived, pre-existing mRNA as in lily (Linet al., 1987) and petunia (Jackson and Linskens,1982), or by regulating enzyme synthesis at thelevel of transcription as in wheat (Bianchettiand Sartirana, 1967), barley (Katayama andSuzuki, 1980), maize (Maugenest et al., 1999), pea(Kuvaeva and Kretovich, 1978), mung bean(Mandal and Biswas, 1970) and lentils (Greineret al., 2005). In addition, there is good evidencefor the activation of pre-existing inactive phytate-degrading enzymes during early stages of germi-nation (Eastwood and Laidman, 1971; Gabardand Jones, 1986). Many alternative ways can besuggested by which enzymes can be reversiblyinactivated: folding; enclosure in, or attachmentto, membranes; association or dissociation ofsubunits; or addition of a section to the polypep-tide chain of the active protein. In such cases,during germination a reversal of these processescould occur.

Gibberellic acid and phosphate may controlphytate degradation during germination, but sofar there is only a small amount of contradictoryinformation available on the effect of both com-pounds on phytate-degrading activity in plantseeds and pollen. In maize, gibberellic acid doesnot significantly alter the accumulation of phy-tate-degrading enzymes during germination(Maugenest et al., 1999), but no informationabout its effect on phytate breakdown is avail-able. An enhancement of phytate degradation bygibberellic acid without any effect on measurablephytate-degrading activity was found in barley(Gabard and Jones, 1986) and wheat (Eastwoodand Laidman, 1971). Gabard and Jones (1986)suggested that gibberellic acid merely increasesthe secretion of phytate-degrading enzymes, butdoes not stimulate their synthesis, thus givingphytate-degrading enzymes access to phytate.Eastwood and Laidman (1971) claimed thatgibberellic acid stimulates phytate breakdownby releasing phosphate, a potent competitiveinhibitor of many phytate-degrading enzymes(Konietzny and Greiner, 2002), from aleuronecells, where most of the cereal phytate-degradingactivity is located. In barley (Katayama andSuzuki, 1980) and lentils (Greiner et al., 2005),gibberellic acid was shown to enhance phytate-degrading activity and phytate degradation dur-

ing germination and it was concluded that thiseffect of gibberellic acid is due at least in part to astimulation of de novo enzyme synthesis partici-pating in phytate breakdown. Centeno et al.(2001) reported an increase in phytate-degradingactivity accompanied by a reduction in phytatebreakdown in rye and barley in the presence ofgibberellic acid, but gave no interpretation forthis apparently contradictory observation. Twomain mechanisms appear to be involved inthe regulation of phytate-degrading activity byphosphate. Most phytate-degrading enzymes arestrongly inhibited by phosphate in vitro (Konietznyand Greiner, 2002), so enzyme activity itselfmay also be controlled by phosphate in vivo(Eastwood and Laidman, 1971; Katayama andSuzuki, 1980). Furthermore, it was concludedthat phosphate also acts at the transcription levelthrough repression of phytate-degrading enzymeexpression (Sartirana and Bianchetti, 1967).

One explanation for the inconsistency of theavailable data on the regulation of phytate-degrading activity in plant seeds and pollenduring germination is the presence of severalmolecular forms of phytate-degrading enzymes ina certain plant (Kuvaeva and Kretovich, 1978;Goel and Sharma, 1979; Baldi et al., 1988;Konietzny et al., 1995; Hamada, 1996; Mauge-nest et al., 1999; Greiner et al., 2000b; Greiner,2002) that are regulated in different waysand may have different physiological functions.Different analytical approaches to determinephytate-degrading activity may also contribute tothe conflicting findings. It was previously shownthat values obtained by extraction methods areconsiderably lower than those obtained by directincubation methods (Greiner and Egli, 2003).

In vivo Function of Phytate-degrading Enzymes

Phytate-degrading enzymes in plant seeds andmicroorganisms occur in multiple forms (Kuvaevaand Kretovich, 1978; Goel and Sharma, 1979;Baldi et al., 1988; Hamada, 1994; Konietzny et al.,1995; Maugenest et al., 1999; Greiner et al.,2000b; Cottrill et al., 2002; Greiner, 2002). Theseforms may exhibit different stereospecificity ofphytate dephosphorylation, be regulated in differ-ent ways, be directed to different localization


within and outside the producing cell, and thusmay have different physiological functions. As theclassification of ‘phytases’ is solely according totheir capability of hydrolysing phytate in vitro,some of these enzymes may not be involved inphytate degradation in vivo but may have com-pletely different functions.

Proposed functions of phytate-degradingenzymes in plants

Phytate-degrading enzymes in higher plants occurpredominantly in grains, seeds and pollen, but littleinformation is available on their specific localiza-tion within these. In cereal seeds, phytate-degrad-ing activity was found to be mainly associated withthe aleurone layer (Gabard and Jones, 1986),whereas in legume seeds it was reported to belocated in the cotyledons (Gibson and Ullah, 1988;Hegeman and Grabau, 2001). Further, the extrac-tion of phytate-degrading activity is stronglyenhanced by the presence of Triton X-100, sug-gesting an association with membrane structures(Scott and Loewus, 1986; Scott, 1991; Greiner andEgli, 2003). The germination-inducible enzymes inparticular are responsible for phytate breakdownduring germination to make phosphate, mineralsand myo-inositol available for plant growth anddevelopment (Greiner et al., 2005).

Some phytate-degrading activity is alsofound in plant roots (Hübel and Beck, 1996; Liet al., 1997; Hayes et al., 1999), in which it mayplay a role in providing the central stele withminerals (Maugenest et al., 1999) or allow plantsto use soil inositol phosphates. However, phytateappears to be only poorly utilized by plants,probably due to a combination of low phytate-degrading activity of roots and the low solubilityof soil phytate (Hayes et al., 2000; see alsoRichardson et al., Chapter 15, this volume). Thus,it was suggested that soil microorganisms coloniz-ing the plant rhizosphere and producing extracel-lular phytate-degrading activity, such as Bacillusand Enterobacter ssp., or membrane-bound phy-tate-degrading enzymes such as those synthesizedby mycorrhizal fungi, could act as plant growthpromoting microorganisms by making phytatephosphate available to the plant (McElhinneyand Mitchell, 1993; Richardson et al., 2001; Idrisset al., 2002).

Possible roles of phytate-degradingenzymes in microbes

The efficient de-repression of phytate-degradingenzyme formation by phosphate starvation inmost microorganisms suggests a possible role forthese enzymes in providing the cell with phos-phate. This is supported by the identification of aphytate-degrading enzyme in the stalk ofCaulobacter crescentus, an aquatic bacterium thatlives in oligotrophic environments where phos-phate limits productivity (Ireland et al., 2002).Phosphate uptake is one of the hypothesizedfunctions of the stalks, which is enhanced whenthe stalks elongate during phosphate limitation.This increase in surface area, as well as the pres-ence of a phytate-degrading enzyme, would allowthe uptake of organic phosphate. With respect tophytate utilization it is also worth mentioningthat cyanobacteria belonging to Rivulariaceaeshowed better growth in phytate when they formhairs compared to non-hair-forming Rivulariaceae(Whitton et al., 1991). The assumption could alsoexplain why, with the exception of sourdoughbacteria, there is no clear evidence for lactic acidbacteria with the ability to degrade phytate.Lactic acid bacteria are adapted to environmentsrich in nutrients and energy where evolutionaryselection pressure would not favour the capabilityto produce a phytate-degrading enzyme.

Phytate-degrading enzymes in S. cerevisiae arepart of the PHO protein family, which is involvedin the scavenging and specific uptake of phos-phate from extracellular sources (Lemire et al.,1985). In E. coli, however, the phytate-degradingenzymes are not under the control of the PHOregulon (Touati et al., 1987). Therefore, they donot appear to have a primary role in phosphateassimilation, even though they probably con-tribute to periplasmatic phosphate levels undercertain conditions. The agp-encoded phosphataseacts primarily as a glucose scavenger, whereas theappA-encoded phosphatase is believed to have arole in stress protection. In E. coli, the primaryresponse to the limitation of a specific nutrient isactivation of a certain set of genes that improvesuptake of the nutrient present in low concentra-tion or allows the utilization of other substancesthat belong to the same class of nutrient. Thesenutrient-specific systems include the cAMP–CRPregulon for the use of alternative carbon sources,the NtrB/NtrC/σ54 regulon that is induced

88 R. Greiner

under nitrogen limitation and the PhoB/PhoRregulon that is induced under phosphorus limita-tion, as well as systems for scavenging low con-centrations of iron and other essential substances(Hengge-Aronis, 1996).

If alternative nutrient sources are present inthe growth medium, the cells continue to growand divide. However, if the environment is totallyexhausted of an essential nutrient, the cells enterinto the stationary phase. In contrast to the spe-cific responses, the nature of the stationary phaseresponse does not appear to be dependent on thelimiting nutrient. Thus, the reaction to nutrientlimitation can be seen as a two-stage process. Ifthe induction of the nutrient-specific responsesremains unsuccessful (i.e. growth cannot beresumed), the stationary phase response is included.This secondary response involves a transition froma metabolic state aimed at maximal growth andcell division to a maintenance metabolism and theinduction of many genes whose function it is toprovide maximal protection against a large varietyof stress conditions.

In contrast to most other microorganisms,anaerobic rumen bacteria are capable of toleratinga high level of phosphate without any negativeimpact on phytate-degrading enzyme formation(Yanke et al., 1998). This unique ability may resultin efficient phytate hydrolysis in the rumen, evenunder the high phosphate levels in the rumen fluidof ruminants fed concentrated feed. However, thisraises the question about the physiological functionof this bacterial phytate-degrading activity if phos-phate limitation is not a problem.

Alternative functions of phytate-degrading enzymes

To provide the cell with phosphate, phytate-degrading enzymes must have access to phytatein the environment. Extracellular phytate-degrading enzymes have been identified inmoulds and yeast, whereas in bacteria theseenzymes are mainly cell-associated. The onlybacteria showing extracellular phytate-degradingactivity are those of the genera Bacillus (Powarand Jagannathan, 1982; Shimizu, 1992; Kerovuoet al., 1998; Kim et al., 1998) and Enterobacter(Yoon et al., 1996). The phytate-degradingenzymes of E. coli have been reported to be

periplasmatic proteins (Greiner et al., 1993).However, phytate is equally well hydrolysedby both disrupted and intact E. coli cells(R. Greiner, 2005 unpublished data). Con-sequently these enzymes appear to have freeaccess in vivo to the substrates present in the sur-rounding medium. Phytate-degrading activity inS. ruminantium and M. multiacidus was found to beassociated with the outer membrane, eventhough phosphate was released from phytate intothe culture fluid by pure cultures (D’Silva et al.,2000). It is not known how bacteria with anapparent lack of extracellular phytate-degradingactivity, such as some Pseudomonas strains, eithergrow in the absence of a readily utilizable phos-phate source or acquire phosphate. As no phos-phate was detected in the growth medium eitherinitially or throughout the growth period(Richardson and Hadobas, 1997), phytate mightbe transported into the bacterial cells. Wang et al.(2004) suggested, for example, that in K. pneumo-niae the gene encoding the phytate-degradingenzyme is co-transcribed from a polycistronicmRNA, which also acts as a template for an inos-itol phosphate transporter.

The role of phytate-degrading enzymes isnot limited to simple degradation functions inmetabolic pathways. In fact, myo-inositol phos-phate phosphatase activities were shown to beinvolved in signal transduction, cell division andmicrobial pathogenesis (Craxton et al., 1997;Zhou et al., 2001). The discovery of myo-inositolfluxes as a result of pathogen infections is anintriguing finding, particularly as phospholipidsignalling plays a critical role in many host cellfunctions, including those affecting cellular sur-vival and regulation of intracellular membranetrafficking (DeVinney et al., 2000). Although sev-eral pathogens trigger myo-inositol phosphatefluxes, different mechanisms appear to be in-volved. Many gram-negative animal and plantpathogens have developed specialized type IIIsecretion systems to translocate bacterial proteinsdirectly into the eukaroytic host cell (DeVinneyet al., 2000). These can influence survival, inter-nalization and replication of the pathogens. Forenteropathogenic E. coli, Helicobacter pylori andSalmonella typhimurium it was proposed that inti-mate contact between the pathogen and the hostcell stimulates host cell phospholipase C activity,resulting in the cleavage of phosphatidylinositol4,5-bisphosphate into the second messengers


D-myo-inositol 1,4,5-trisphosphate and diacylglyc-erol (Ruschkowski et al., 1992; Foubister et al.,1994). For Listeria monocytogenes, a secreted phos-phatidylinositol-specific phospholipase C (PI-PLC)was suggested to be an essential determinant oftheir pathogenesis (Camilli et al., 1991).

A number of further gram-positive humanpathogens secrete PI-PLCs, including Staphylo-coccus aureus, Clostridium novyi, B. cereus andB. anthracis (Low, 1989). Salmonella dublin wasreported to modulate host cell signalling path-ways by translocating at least two proteins, SopBand SopE (Zhou et al., 2001). SopB exhibits myo-inositol phosphate phosphatase activity andspecifically dephosphorylates D-myo-inositol1,3,4,5,6-pentakisphosphate to D-myo-inositol1,4,5,6-tetrakisphosphate. A phosphatase capableof catalysing the same reaction was also found inother Enterobacteriaceae family members, such as E.coli (Cottrill et al., 2002), P. agglomerans (Greiner,2004) and E. cloacae (R. Greiner, 2004 unpub-lished data). Due to the unique myo-inositol phos-phate phosphatase activity of these enzymes,Cottrill et al. (2002) raised the question of a rolefor them in microbial pathogenesis or cellularmyo-inositol phosphate metabolism. However, incontrast to SopB (Norris et al., 1998), these threeenzymes also hydrolyse phytate in vitro. SopE hasno inherent phosphatase activity, but is proposedto activate an endogenous cellular myo-inositolphosphate phosphatase (Zhou et al., 2001). Inaddition, both SopB and SopE stimulate cellularresponses that lead to host cell phospholipase Cactivation.

Xanthomonas oryzae pv. oryzae, an importantrice pathogen, has been suggested to require aBacillus-like phytate-degrading enzyme for opti-mal virulence (Chatterjee et al., 2003).Homologues of this gene are also present in thegenomes of X. campestris pv. campestris andX. axonopodis pv. citri. The enzyme is secreted andmay interfere with myo-inositol phosphate-basedsignalling processes in plants. Differencesbetween this Xanthomonas protein and the Bacillusphytate-degrading enzyme, specifically inresidues involved in the binding of the phosphateresidue attached to the D-2 position of the phy-tate molecule (Chatterjee et al., 2003), may resultin an inability to bind phytate and other myo-inositol phosphates phosphorylated at the D-2position. Therefore, this Xanthomonas enzyme mayexhibit a myo-inositol phosphate phosphatase

activity that emulates the virulence-associatedSopB phosphatase from S. dublin.

Position D-2 is the only one in axial orienta-tion, and all myo-inositol phosphate intermediatesidentified so far during enzymatic phytate break-down are phosphorylated in this position.However, with the exception of phytate itself,intracellular myo-inositol phosphates are alwaysdephosphorylated at the D-2 position. The cellmay therefore discriminate between intra- andextracellular-generated myo-inositol phosphatesby the phosphorylation status of the D-2 positionof the myo-inositol ring. Further, a considerablebody of evidence supports the hypothesis that thephosphorylation status of the D-2 position is usedto independently control the synthesis and break-down of phytate in the plant kingdom. For exam-ple, this was concluded from the myo-inositolphosphates identified in a development stage ofthe plant Spirodela polyrhiza L., which is associatedwith massive accumulation of phytate (Brearlyand Hanke, 1996a,b). None of these myo-inositolphosphates was phosphorylated at the D-2 posi-tion of the myo-inositol ring.

Conclusions and Future ResearchDirections

Until now, research on phytate-degradingenzymes has focused almost exclusively on mak-ing enzymes available that are suitable for use asanimal feed additives. The in vivo function of phy-tate-degrading enzymes has therefore received lit-tle attention. So far, only the germination-inducible phytate-degrading enzymes of plantseeds could be called phytases. Their action uponphytate is considered likely, because the break-down of phytate during germination makes phos-phate, minerals and myo-inositol available forplant growth and development. As formation ofextracellular phytate-degrading enzymes inmoulds and yeast is triggered by phosphate star-vation, these enzymes hydrolyse organic phos-phates, including phytate, to provide the cell withphosphate from extracellular sources. Theseenzymes are therefore non-specific phosphatasesthat exhibit phytate-degrading activity. Thein vivo function of other phytate-degradingenzymes is mainly speculative. In addition to pro-viding the cell with phosphate as mentioned

90 R. Greiner

above, a role in stress response or bacterialpathogenesis has been assumed.

Information on the regulation of phytate-degrading enzyme formation might shed light onthe in vivo role of phytate-degrading enzymes.Extensive studies have so far been performedonly for E. coli and S. cerevisiae, although someinformation on the regulation of phytate-degrad-ing enzyme formation is also available for R. terri-gena and germinating cereal and legume seeds.The majority of studies, however, were per-formed on a trial-and-error basis to optimizemedium composition with respect to phytate-degrading enzyme yields. Micro-arrays should beused to learn more about the regulation of phy-tate-degrading enzyme formation and their in vivofunctions, in addition to classical biochemical andmolecular approaches. Running transcriptomeprojects may provide useful information on theexpression of genes encoding phytate-degradingenzymes. There are already large data-sets, e.g.for yeast (http://www.transcriptome.ens.fr/ymgv)and Arabidopsis (https://www.genevestigator.

ethz.ch), online which could be mined for theinformation they contain on the environmen-tal and biotic factors that contribute to expres-sion of genes encoding phytate-degradingenzymes.

In addition to understanding the regulationof phytate-degrading enzyme formation and theirin vivo function, a number of other importantquestions remain to be answered. Only a rela-tively small number of organisms have been iden-tified as being able to utilize phytate as the solephosphate or phosphate and carbon source, butthe prevalence of this ability among microbesremains unknown. More information is requiredon the ecology of phytate-utilizing bacteria,where they occur, and the site of phytate dephos-phorylation. Furthermore, it has not been clearlyestablished that only extracellular phytate-degrading enzymes make phytate-phosphateavailable to the microbial cell. It thereforeremains possible that phytate is transported intactacross the microbial cell wall to be dephosphory-lated within the cell.


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96 R. Greiner

7 Phytases: Attributes, CatalyticMechanisms and Applications

Edward J. Mullaney and Abul H.J. UllahUnited States Department of Agriculture–Agricultural Research Service,

Southern Regional Research Center, 1100 Robert E. Lee Blvd, New Orleans,LA 70124, USA


Discovered more than a century ago (Posternak,1903), myo-inositol hexakisphosphate is a ubiqui-tous constituent in cereals and grains, where itexists predominantly in salt form (phytate). Thismeans that it represents an immense reservoir ofphosphorus that can be potentially utilized byplants, microorganisms and animals. Lott et al.(2000) estimated that 51 million t of phytate issequestered annually in commercially producedcrop seeds and fruits. The amount of phosphorusin phytate is therefore equivalent to approxi-mately two-thirds of the phosphorus utilizedeach year through the application of mineralfertilizers to agricultural land. Phosphorus isan essential component of DNA, adenosine 5′-triphosphate and other compounds necessaryfor life (Abelson, 1999), so the liberation of inor-ganic phosphate covalently bound to phytateis essential in numerous organisms. The firstphytate-degrading enzyme was reported as earlyas 1907 (Suzuki et al., 1907) and research contin-ues to the present day.

A major impetus for phytase researchresulted from the poultry industry switching fromfishmeal and other more expensive proteinsources to low-cost plant protein such as soybeanmeal (Rumsey, 1993). Poultry and other animalswith simple stomachs lack a digestive phytase, soresearch to identify a cost-effective phytase thatcould be added to their diet intensified (Wodzinski

and Ullah, 1996). Today, a number of enzymesthat can initiate the cleavage of myo-inositol hexa-kisphosphate are known to exist in a range oforganisms (Konietzny and Greiner, 2002; Simonand Igbasan, 2002; Lei and Porres, 2003;Mullaney and Ullah, 2003; Oh et al., 2004;Haefner et al., 2005; see Hill and Richardson,Chapter 5, and Greiner, Chapter 6, this volume).The detailed characterization of some of theseenzymes has revealed that nature did not developa single catalytic mechanism to cleave phosphategroups from myo-inositol hexakisphosphate inthese diverse organisms. In fact, the exact num-ber of catalytic mechanisms that nature hasevolved for this purpose is not known. However,it is now clear that different strategies havebeen adopted to accommodate the physical struc-ture of myo-inositol hexakisphosphate with itssix negatively charged phosphate groups as asubstrate.

The recognition that not all phytases arestructurally similar or share a common active sitehas already yielded an initial classification systembased primarily on catalytic mechanism (Ohet al., 2004). However, a taxonomic system needsto be devised that can accommodate new types ofphytases with novel catalytic mechanisms. Theincreasing number of phytate-degrading enzymesnow offers scientific investigators the potential toselect the catalytic features most amenable to

their research objectives. Coupled with thepotential to engineer certain enzyme characteris-tics, this offers the potential for significantachievements in this and related fields ofresearch.

Today, four distinctly different classes ofphosphatase enzymes are known to display phy-tase activity:

1. Histidine acid phosphatase (HAP)2. β-Propeller phytase (BPP)3. Cysteine phosphatase (CP)4. Purple acid phosphatase (PAP)

The representatives of each of these classes havedifferent catalytic mechanisms and other uniquefeatures that allow them to effectively utilize myo-inositol hexakisphosphate as a substrate at vari-ous pH values. The fact that each is different alsobroadens the number of enzymes that may beapplied to the search for developing an enhancedphytase for current and future applications ofsuch enzymes.

Histidine Acid Phosphatase

HAP is a large class of enzymes with representa-tives in animals, plants and microorganisms(Wodzinski and Ullah, 1996; Mullaney et al.,2000; Konietzny and Greiner, 2002; Lei andPorres, 2003). A shared characteristic is a com-mon catalytic mechanism. The amino acid

sequences that compose the active site have beenidentified and contain an N-terminal motifRHGXRXP and a C-terminal motif HD. Whenproperly folded, these distant sequences convergeto form a single catalytic centre, which initiates atwo-step reaction that hydrolyses phosphomo-noesters (Ullah et al., 1991; van Etten et al., 1991).In this reaction, the histidine residue in the con-served motif of the N-terminal region serves as anucleophile in the formation of a covalent phos-pho-histidine intermediate (Ostanin et al., 1992;Lindqvist et al., 1994; Oh et al., 2004). The aspar-tic acid residue of the C-terminal HD then func-tions as a proton donor to the oxygen atom of thescissile phosphomonoester bond (Lindqvist et al.,1994; Porvari et al., 1994).

The N-terminal active-site motif and theadjacent amino acids found in a wide array ofHAPs are presented in Fig. 7.1. It should benoted that the exact sequence is even conservedin the prokaryotic example, Escherichia coli phy-tase, shown in this figure. It must also be notedthat while these enzymes share a common cat-alytic site, they do not share an equal ability todegrade myo-inositol hexakisphosphate. Both themouse and fruit fly multiple inositol polyphos-phate phosphatase (MIPP) in Fig. 7.1 represent anumber of other HAPs that are not effective phy-tases. Also, the fact that maize phytase displaysonly 60% homology with the HAP consensusmotif (Maugenest et al., 1999), yet is still a phytase,indicates that the ability to utilize myo-inositol

98 E.J. Mullaney and A.H.J. Ullah

HAP consensus sequence

Aspergillus niger NRRL 3135 PhyA

Aspergillus terreus PhyA

Aspergillus nidulans PhyA

Escherichia coli phytase (appA gene product)

Mus musculus, multiple inositol polyphosphate phosphatase (MIPP)

Drosophila melanogaster MIPP

Maize Phyt I

RHGXRXP

QVLSRHGARYPTSK

QVLARHGARSPTDS

QVLSRHGARYPTES

VIVSRHGVRAPTKA

VALIRHGTRYPTTK

MWIFRHGDRTPKKS

ELVRRHQLRLGYGS

Fig. 7.1. The conserved N-terminal active site and flanking amino acid sequence in representativehistidine acid phosphatases from microorganisms, animals and plants. All enzymes shown, exceptmouse and Drosophila MIPP (multiple inositol polyphosphate phosphatase), display phytase activity.

hexakisphosphate as a substrate depends on morethan a single catalytic feature. In recognizing thefact that all HAPs are not phytases, Oh et al.(2004) advanced the term ‘histidine acid phytase’(HAPhy) to denote HAPs that can accommodatemyo-inositol hexakisphosphate as a substrate.

Both prokaryotic and eukaryotic HAPhyenzymes are known and share little sequencehomology other than the conserved active-sitemotif. The E. coli phytase is the best-characterizedprokaryotic HAPhy (Greiner et al., 1993) and athree-dimensional molecular model of its struc-ture is available (Lim et al., 2000). This enzymehas been advanced for use as an animal feed addi-tive. The successful expression of the E. coli phy-tase gene in the salivary glands of mice (Golovanet al., 2001a) and swine (Golovan et al., 2001b) wasreported recently. The swine, termed theEnviropig™, secretes phytase in its saliva to breakdown phytate in its feed and produce low-phos-phorus manure.

Substrate specificity site

In eukaryotes, HAPhys have been cloned inmaize and a number of fungal isolates. The mostwidely studied fungal phytases are from Aspergillusniger and A. fumigatus. The crystal structure ofboth of these enzymes has been derived anddeposited in the National Center for Bio-technology Information (NCBI) as 1IHP and1SK8, respectively. Structural characterization(Kostrewa et al., 1999; Liu et al., 2004; Xianget al., 2004) and catalytic studies (Wyss et al.,1999a) have identified a new site in the enzymethat facilitates its interaction with different sub-strates. Kostrewa et al. (1999) identified a region,the substrate specificity site, of the A. niger PhyAmolecule that encircles the active site of theenzyme and functions as a ‘gatekeeper’. Thesame site allows for the interaction of the cat-alytic site and the highly negatively charged myo-inositol hexakisphosphate molecule. In thesubstrate specificity site of A. niger NRRL 3135there are two acidic and four basic amino acidresidues, E228, D262, K91, K94, K300 andK301 (Kostrewa et al., 1999; Mullaney et al.,2000). This means that at pH 2.5 the four basicamino acids – K91, K94, K300 and K301 – areall positively charged and would attract myo-

inositol hexakisphosphate. When the pH is raisedto 5.0, the local electrostatic field of the substratespecificity site still remains attractive for myo-inositol hexakisphosphate.

Wyss et al. (1999a) had previously observedthat despite the catalytic centres for all the knownmicrobial HAPhys being identical, they could bedivided into two classes based on substrate speci-ficity. One class has broad substrate specificity buta low specific activity for myo-inositol hexakisphos-phate, while the second class has narrow substratespecificity and a high specific activity for myo-inos-itol hexakisphosphate. An examination of theamino acids composing the substrate specificitysite of the fungal HAPhys in this study revealed acorrelation between the amino acid residue 300and the enzyme’s level of specific activity for myo-inositol hexakisphosphate (Mullaney et al., 2002).This study also revealed that residue 301 wasstrongly conserved as lysine (K), while residue 300varied considerably. The HAPhys cited in Wysset al. (1999a) with high specific activity for myo-inositol hexakisphosphate have either a basic oracidic amino acid residue at 300, while the phy-tases with low specific activity have a neutralamino acid at that position. Subsequent site-directed mutagenesis at residue 300 in A. nigerNRRL 3135 PhyA established the importance ofthe lysine residue at that site and the enzyme’shigh specific activity for myo-inositol hexakisphos-phate (Mullaney et al., 2002). It should be notedthat the replacement of amino acids that are notpart of the substrate specificity site has also beenreported to enhance the catalytic properties of A.fumigatus phytase (Tomschy et al., 2000).

Site-directed mutagenesis studies have linkedsome of the amino acid residues in the substratespecificity site of A. niger NRRL 3135 to its uniquepH, with two optima at 2.5 and 5.0 (Ullah andGibson, 1987). Explanations for this phenomenonrange from dismissal as an artefact (Berka et al.,1998) to possible buffer effects (Lehmann et al.,2000). The selection of an appropriate buffer is ofcourse critical in determining the true specificactivity any enzyme displays for myo-inositol hexa-kisphosphate at various pH values. Recent studiesin both A. niger NRRL 3135 (Mullaney et al.,2002) and A. fumigatus ATCC 13073 PhyA(Tomschy et al., 2002) have established that theamino acid residues in the substrate specificity sitegive rise to this unique two-pH optima profile.

Phytases: Attributes, Catalytic Mechanisms and Applications 99

Differences in the substrate specificity site ofa second extracellular A. niger phytase, PhyB, canalso explain why it has both a different pH opti-mum and substrate specificity range than A. nigerPhyA. PhyB has only been reported in the iso-lates of A. niger. Although the active form of PhyAis a monomer, PhyB was reported to be either adimer (Ullah and Cummins, 1987) or a tetramer(Kostrewa et al., 1999). The individual moleculesare electrostatically bound together at the N-terminal region. PhyB displays an optimumpH at 2.5, but unlike PhyA it cannot hydrolysemyo-inositol hexakisphosphate at pH 5.0. That iswhy researchers named it pH 2.5 optimum acidphosphatase when first characterized (Ullah andCummins, 1987). Subsequent investigationsrevealed that it could hydrolyse myo-inositol hexa-kisphosphate at pH 2.5 (Ullah and Phillippy,1994). Both PhyB and PhyA share identicalactive-site characteristics of HAPs, but their sub-strate specificity sites are remarkably different.Kostrewa et al. (1999) identified the substratespecificity site of PhyB to be composed of onlytwo acidic amino acids, D75 and E272. Thismeans that at pH 5.0 the acidic amino acidswould be negatively charged, while at pH 2.5they would be uncharged. All negatively chargedsubstrates, including myo-inositol hexakisphos-phate, would therefore be repelled at the higherpH, but not at the lower. This also explains thesubstrate specificity difference between PhyA andPhyB. The latter can accept a broader variety ofphosphomonoesters, because its substrate speci-ficity site has a more neutral electrostatic field,whereas A. niger’s PhyA has a highly positive elec-trostatic field at its substrate specificity site and isthus optimized for the binding of negativelycharged myo-inositol hexakisphosphate.

All the recent findings about the substratespecificity site of fungal HAPs suggest that it hasa significant role in determining how effectivelythe enzyme can hydrolyse myo-inositol hexak-isphosphate. By occupying positions adjacent tothe catalytic domain, the amino acids in the sub-strate specificity site function as gatekeepers indetermining the ease with which any substratecan pass and interact with the active-site residues.Research is also showing that techniques such assite-directed mutagenesis of a cloned phytasegene can be employed to alter the composition ofthe enzyme’s substrate specificity site and thusalter both its pH profile and substrate selectivity.

Glycosylation

The desire to obtain a commercially viable phy-tase has focused most research on the isolation ofextracellular or secreted enzymes. Consequently,all the fungal phytases that have been charac-terized thus far are secreted glycoproteins.Glycosylation, the process that adds polysaccha-rides to proteins, is generally thought to conferstability and assist in the correct folding of theenzyme. The SDS–PAGE profile of purifiedPhyA provided the first clue that it was heavilyglycosylated (Ullah and Gibson, 1987). Sugaranalysis of purified phytase indicated the pres-ence of N-glycosidic linkage of high mannosesugar chain to the asparagines (N) (Ullah, 1988).This conclusion was bolstered during chemicalsequencing of phytase (Ullah and Dischinger,1993), which revealed several blank residueslinked to the presence of a consensus sequence,or sequon, NXS in the PhyA sequence (Apweileret al., 1999). The presence of extensive glycosy-lation on fungal phytase impeded its crystalliza-tion for structural studies. It was not untilGrueninger-Leitch et al. (1996) developed arecombinant fusion protein glycosidase expressedin E. coli that it became feasible to obtain thehigh-quality protein crystals that were required todetermine a proper three-dimensional X-raystructure of glycoproteins. A. niger PhyA was oneof the first enzymes deglycosylated by this tech-nique. All ten N-glycosylation sites of A. nigerNRRL 3135 PhyA are indicated in Fig. 7.2.

Patterns of glycosylation vary in phytasewhen expressed in other fungal expression sys-tems and transgenic plants (Ullah et al., 1999,2002; Wyss et al., 1999b). However, the activity ismaintained, and PhyA has been successfullyexpressed in several fungal expression systemsand a number of plant species (Mullaney et al.,2000). Differences in the glycosylation pathwayin fungi and animals may explain the low successrate in the expression of any fungal phytase inanimals (Bretthauer, 2003). To date, A. niger phy Ahas been reported to be expressed in only twoanimals: a fish, the Japanese medaka, Oryziaslatipes (Hostetler et al., 2003), and silkworm,Bombyx mori (Wang et al., 2003). However, highersuccess rates have been obtained with the ungly-cosylated prokaryotic E. coli phytase, which hasbeen expressed in both mice and swine (Golovanet al., 2001a,b).


Disulphide bridges

Although not directly involved in the catalyticfunction of HAPhy, disulphide bridges performan important role in maintaining the properthree-dimensional structure to allow for catalyticactivity in phytase (Ullah and Mullaney, 1996;Kostrewa et al., 1997; Wang et al., 2004). Thethree-dimensional models of A. niger and A. fumi-gatus PhyA show that all ten cysteine residuespresent in each enzyme are involved in the for-mation of five disulphide bridges. An analysis ofthe deduced amino acid sequence of a numberof other fungal phytases that have not been soextensively characterized revealed a pattern ofconservation of the cysteines necessary for theformation of these disulphide bridges (Mullaneyand Ullah, 2005).

A shared eight-cysteine motif (8CM) iswidely conserved in all the fungal HAPhys sur-veyed. The reason for the conservation of this8CM appears to parallel the recent discovery ofanother eight-cysteine motif in a number of plantproteins (Jose-Estanyol et al., 2004). In none ofthese nearly 500 plant polypeptides were any ofthe cysteines involved in the functional catalyticmechanism of the molecules. Instead, they wereconserved to form a network of disulphide bondsthat allow for the convergence of sequences nec-

essary for the proper molecular architecturerequired for folding and specific function of theproteins.

In fungal HAPhys, the conservation of the8CM sequence was 100%, while the overallhomology for the sequences examined rangedbetween 23% and 66%. The survey includedamino acid sequences from basidiomycete(Lassen et al., 2001), unicellular and filamentousascomycete HAPhys (Mullaney and Ullah, 2005).This study also reported that two extra cysteines,which form a fifth disulphide bridge in the N-terminal region of A. niger PhyA, are conserved inall filamentous ascomycete HAPhys. This sug-gests that the higher stability found in phytasesfrom some Aspergillus spp. may in some mannerbe correlated with this extra disulphide bridge.

In E. coli phytase, all eight cysteines areinvolved in four disulphide bonds (Lim et al.,2000). However, in this phytase, significantlyenhanced activity was achieved when one disul-phide bridge was abolished. This was accom-plished by site-directed mutagenesis, whichreplaced a cysteine with another amino acidresidue (Rodriguez et al., 2000). It was sug-gested that the removal of this disulphide bondcould modulate the domain flexibility andthereby increase the catalytic efficiency of theenzyme.


# #MGVSAVLLPLYLLSGVTSGLAVPASRNQSSCDTVDQGYQCFSETSHLWGQYAPFFSLANE 60

# #SVISPEVPAGCRVTFAQVLSRHGARYPTDSKGKKYSALIEEIQQNATTFDGKYAFLKTYN 120

YSLGADDLTPFGEQELVNSGIKFYQRYESLTRNIVPFIRSSGSSRVIASGKKFIEGFQST 180

# #KLKDPRAQPGQSSPKIDVVISEASSSNNTLDPGTCTVFEDSELADTVEANFTATFVPSIR 240

QRLENDLSGVTLTDTEVTYLMDMCSFDTISTSTVDTKLSPFCDLFTHDEWINYDYLQSLK 300

# #KYYGHGAGNPLGPTQGVGYANELIARLTHSPVHDDTSSNHTLDSSPATFPLNSTLYADFS 360

# #HDNGIISILFALGLYNGTKPLSTTTVENITQTDGFSSAWTVPFASRLYVEMMQCQAEQEP 420

LVRVLVNDRVVPLHGCPVDALGRCTRDSFVRGLSFARSGGDWAECFA

Fig. 7.2. The amino acid sequence of Aspergillus niger NRRL 3135 PhyA. All ten N-glycosylation sitesare indicated by #.

Applications

Today, the major application for HAPhys is inthe hydrolysis of phytate in cereal grains of ani-mal feed. Numerous feed trials involving poultry,swine and aquaculture have established theefficacy of commercially produced phytase toincrease the utilization of phytate while reducingphosphorus concentrations in manure (Kornegay,1996; Wodzinski and Ullah, 1996; Mullaneyet al., 2000). Phytase is now extensively employedworldwide for this application, which is discussedin detail elsewhere in this volume (Lei andPorres, Chapter 9).

Food applications include the production ofphytate-free soybean milk, low-phytin bread(Dvorakova, 1998; Brinch-Pedersen et al., 2003;Drakakaki et al., 2005) and the production oftransgenic grains expressing fungal phytase(Lucca et al., 2001). One novel application wasthe incorporation of vanadium into the active siteof A. niger PhyA to produce a cost-effective semi-synthetic peroxidase (van de Velde et al., 2000).A limited market also exists for the use of HAPhyin the production of myo-inositol phosphates.Greiner and Konietzny (1996) demonstrated thata packed-bed bioreactor containing covalentlyattached E. coli PhyA could cost-effectively pro-duce specific isomers of the lower myo-inositolphosphate esters.

HAPhys have been successfully expressed incrop plants demonstrating the potential for ‘bio-farming’. The goal in these studies was to provethe feasibility of utilizing transgenic crop plants toproduce several enzymes that are currently man-ufactured at conventional fermentation facilities.To document that this was a cost-effective meansto produce bulk enzymes, A. niger phy A was suc-cessfully expressed in the fodder crop Medicagosativa (lucerne) (Ullah et al., 2002).

Future applications of HAPhys extend to thepotential development of plant cultivars thatrequire less phosphorus fertilizer. This researchhas been prompted by the realization that globalphosphate reserves are limited and that futuregenerations may face a ‘phosphate crisis’(Abelson, 1999). To respond to this challenge, theA. niger phy A gene has been expressed in the rootsof a plant, Arabidopsis (Mudge et al., 2003). Thesetransgenic plants can grow on phytate as the solephosphorus source, which opens up the possibil-ity that crop plants developed from this tech-

nique would require less phosphorus fertilizersand thus slow the depletion of the known supplyof phosphorus. To determine the feasibility ofthis in an agricultural crop, George et al. (2004)engineered a transgenic Trifolium subterraneum L.that exudes an extracellular A. niger PhyA from itsroots. This technology is discussed elsewhere inthis volume (Richardson et al., Chapter 15).

b-Propeller Phytase

Unlike HAPhys, which are members of a well-studied class of enzymes, β-propeller phytase(BPPhy) represents an entirely new class ofenzymes and exhibits no homology to any knownphosphatases (Kerovuo et al., 1998; Kim et al.,1998a,b; Ha et al., 2000). The name was adoptedfor this group of enzymes based on their molecu-lar structure, which consists mainly of β-propellersheets and resembles a six-bladed propeller (Haet al., 2000; Shin et al., 2001). Before this, it wastermed PhyC (Kerovuo et al., 1998) and TS-Phy(Ha et al., 1999) and has been subsequentlylabelled PhyD (alkaline phytase) (Oh et al., 2004),PhyL (Tye et al., 2002) and PhyA (Chatterjeeet al., 2003).

Initially, BPPhys were reported from Bacillusand related bacterial species that require calciumions for both catalytic activity and thermosta-bility. The calcium facilitates the binding ofmyo-inositol hexakisphosphate by generating afavourable electrostatic environment in the sub-strate-binding domain of the biocatalyst. Kineticstudies established that BPPhys could hydrolysecalcium-phytate between pH 7.0 and 8.0 (Ohet al., 2001). The main components involved inthe catalytic mechanism of BPPhys to hydrolysemyo-inositol hexakisphosphate include a ‘cleavagesite’ and an ‘affinity site’ (Shin et al., 2001). In thismodel, it is necessary for two adjacent phosphategroups to occupy both the cleavage and affinitysites. The phosphate bound to the affinity sitefacilitates the cleavage of flanking phosphate bythe cleavage site. The enzyme prefers hydrolysisof every second phosphate and has a reducedaffinity for any substrate that cannot accommo-date this stringent requirement. This explainswhy BPPhys alternately remove phosphate groupswith the end product being myo-inositol trisphos-phate. Further degradation then occurs slowly


because a neighbouring phosphate group is lack-ing and the enzyme is increasingly susceptible toproduct inhibition.

Based on its narrow substrate range, arequirement for calcium for catalytic activity, andmyo-inositol trisphosphate being the predominantproduct from myo-inositol hexakisphosphatehydrolysis, Oh et al. (2004) proposed that alkalinephytases from plants share a similar catalyticmechanism with BPPhys. For this reason, severalalkaline plant phytases were grouped in the sameclass with BPPhys. Calcium is known to enhancethe activity of several plant alkaline phytases,among others from lily (Lilium longiflorum) pollen(Scott and Loewus, 1986) and a number oflegumes (Scott, 1991). Unfortunately, none of thegenes thus far have been cloned and no sequencedata exist to confirm that they indeed are BPPhys.However, it should be noted that a recent studyquestions this classification by reporting the pres-ence of the HAP amino acid motif in lily pollenphytase (Mehta and Murthy, 2005).

Another similarity has also been notedbetween BPPhys and pyrophosphatases, whichhydrolyse inorganic pyrophosphate. β-Propellerphytases share no homology with any otherknown classes of phosphatases, which ledHamelryck (2003) to employ a ‘multidimensionalindex tree’ method to analyse side-chain patternsfound in different classes of enzymes. The resultssuggested that BPPhys and pyrophosphatasesshare some common structural features, includ-ing a cleavage site where the nucleophilic attackby a water molecule transpires, and an affinitysite that binds a second phosphate group (Shinet al., 2001).

Although BPPhys and pyrophosphatasesshare a similar catalytic mechanism, the molecu-lar architecture displayed in BPPhys is also foundin a number of other proteins. None of theseother enzymes are phytases, but several oftheir β-propeller domains are associated withpathogenesis in various diseases ranging fromAlzheimer’s and arthritis to microbial infections(Pons et al., 2003). It is interesting to note that theonly phytase associated with pathogenesis to dateis a BPPhy. It was reported that Xanthomonasoryzae, a plant pathogen of rice, secretes a six-bladed β-propeller protein, PhyA, which isrequired for optimum virulence in its host(Chatterjee et al., 2003). Characterization of thisprotein revealed conservation of active-site

residues previously identified in Bacillus phytases.Isolates of X. oryzae having mutant phy A genesdisplay a reduced virulence in their host plants.This is the first study to suggest that virulence ina plant pathogen is due at least in part to theability of the bacteria to utilize myo-inositol hexa-kisphosphate in the plant host as a source ofphosphate. Bacterial leaf blight caused byX. oryzae is a major rice disease and this raisesthe possibility that other bacteria and fungalpathogens of plants have evolved a similar or dif-ferent phytase to access myo-inositol hexakisphos-phate in their host. The naming of this BPPhy,also as PhyA, creates the potential for confusionwith the HAPhy PhyA and reinforces the need toupdate the phytase nomenclature.

β-Propeller phytases have been advocatedfor several applications. Their heat tolerance(Kim et al., 1998a) means that they would with-stand feed-pelleting, which has made them candi-dates for use as an animal feed additive, bothalone and with HAPhy (Park et al., 1999).Research to advance this has resulted in its suc-cessful expression in E. coli to increase its cost-effectiveness (Kim et al., 1998b). Bacillus phytasestimulated the growth of maize seedlings underconditions of phosphate limitation but not in thepresence of myo-inositol hexakisphosphate (Idrisset al., 2002). Bacillus subtilis phytase has also beenexpressed in the cytoplasm of transgenic tobacco(Yip et al., 2003). Results indicated that a shift inthe equilibrium of the inositol phosphate biosyn-thesis pathway occurred, which improved plantperformance under phosphate starvation.Although these studies offer new strategies foranimal feed supplements and possible tools forraising productivity in agriculture, no commercialapplications of BPPhys are currently available.

Cysteine Phosphatase

Another class of phytase has been reported froman anaerobic ruminal bacterium, Selenomonasruminantium. It had long been suspected that thereason ruminants could utilize phytate and mono-gastric animals could not was due in part to thepresence of certain microorganisms in the rumen.A survey of anaerobic rumen bacteria revealedphytase activity in one isolate, S. ruminantium(Yanke et al., 1999). Initial characterization


established that the enzyme was monomeric, ~46kDa in size, had an optimal pH range of 4.0–5.5,an optimal temperature of 50–55ºC and was inhibited by cations of iron and several other met-als. The gene encoding this phytase has now beencloned and its product extensively analysed (Chuet al., 2004). Its crystal structure reveals that it isneither an HAPhy nor a BPPhy. Instead, its struc-ture and proposed catalytic mechanism suggestthat it is a member of the (CP) superfamily. Itshares the active-site motif HCXXGXXR(T/S)and other substantial similarities with protein tyro-sine phosphatase (PTP), a member of the CPgroup. The active site forms a loop that functionsas a substrate-binding pocket unique to PTPs. Thedepth of this pocket is important because itappears to determine the substrate specificity(Denu and Dixon, 1998). Consistent with thismodel, the S. ruminantium phytase, the cysteine phy-tase (CPhy), has a wider and deeper pocket thanPTP and is thus able to accommodate the fullyphosphorylated inositol group of myo-inositol hexa-kisphosphate (Chu et al., 2004).

The initial binding of myo-inositol hexak-isphosphate to the CPhy active-site pocket isfacilitated by the negatively charged substrate.The hydrolysis of phosphate groups proceedssequentially with the end product being myo-inositol 2-monophosphate (Chu et al., 2004). Theinhibitory effect of iron and other metal cations(copper, zinc and mercury) was attributed totheir ability to complex with myo-inositol hexak-isphosphate, but the stimulatory effect of leadcations remains unexplained (Yanke et al., 1999).The source of this enzyme, a ruminal bacterium,suggests that it may have evolved many attributesthat would lead to its adoption as an animal feedadditive. However, at this time no feed trial studyor any other commercial applications of CPhyare available.

Purple Acid Phosphatase

The PAPs have representatives in plants, mam-mals, fungi and bacteria (Schenk et al., 2000). Likeother metalloenzymes, the active site of PAPsrequires one or more metal ions for activity. PAPsshare a pattern of five common consensus motifs(DxG/GDx2Y/GNH(E,D)/Vx2H/GHxH) con-

taining seven residues capable of forming metalligands (Schenk et al., 2000). The first binuclearmetal-containing hydrolase identified as a phytasewas reported in the cotyledons of a germinatingsoybean (Glycine max L. Merr.) seedling (Hegemanand Grabau, 2001). The gene encoding GmPhyhas been cloned and its product characterized.However, unlike HAPhys, BPPhys and CPhys, noX-ray crystallography study has been performedon PAP phytase (PAPhy) and its three-dimen-sional structure is unknown.

An A. niger PAP (Apase6) was isolated andcloned (Ullah and Cummins, 1988; Mullaneyet al., 1995). It does not effectively utilize myo-inositol hexakisphosphate as a substrate andthere has been no commercial interest in this acidphosphatase. Comparison of the active sites ofthis A. niger PAP and the soybean PAPhys(Fig. 7.3) indicates that they both contain theconserved active-site motif (Mullaney and Ullah,1998).

When compared to A. niger PhyA, the spe-cific activity for myo-inositol hexakisphosphate ofGmPhy is low. The lower catalytic activity ofGmPhy may be advantageous during germina-tion because the process requires a steady hydrol-ysis of myo-inositol hexakisphosphate over thisentire period (Mullaney and Ullah, 2003). Chieraet al. (2004) expressed the GmPhy gene in the soy-bean seed during development, a time when it isnot normally expressed. Linking the GmPhy geneto an embryo-specific promoter, β-conglycinin,ectopic expression was achieved and a lower levelof phytate occurred in the transformed seeds.A reduction in phytate of up to 25% wasachieved. This offers another potential strategyfor the development of plant cultivars with lowerphytate levels in their seed.

Since the reporting of GmPhy, putativePAPhys have been reported in the rice (Oryza sativaL.) genome and cloned from a legume, barrelmedic (Medicago. truncatula Gaertn) (Xiao et al., 2005).The latter example revealed high sequence similar-ity to GmPhy, but unlike the soybean phytase, itdoes not have all five conserved blocks of the aminoacids capable of forming metal ligands that arecharacteristic of PAP (Xiao et al., 2005). Only lim-ited characterization of M. truncatula PAPhy is cur-rently available, but it has been engineered to beunder the control of a root-specific promoter,MtPT1, and expressed in Arabidopsis. The trans-


genic phytase was then secreted into the plant’s rhi-zosphere. This permitted the transgenic Arabidopsisto grow when phytate was the sole source of phos-phorus, as reported earlier with the A. niger HAPhygene expressed in Arabidopsis (Richardson et al.,2001). Xiao et al. (2005) suggested that while bothgenes could be employed to improve phosphateacquisition in crops, a plant phytase gene couldhave advantages over a fungal gene with regard toregulatory and biosafety concerns. It was also sug-gested that it would be worth investigating if thesimultaneous expression of the MtPHY1 and a fun-gal phytase gene in the roots of transgenic plantsmight benefit phosphorus uptake. Although noPAPhys are currently being marketed, they havebeen the subject of several research studies.

A Revised Nomenclature forthe Phytases

Many researchers in the field agree that theexpanding interest in phytase over the lastdecade has created the need to consider a revisednomenclature that more accurately describes thecatalytic mechanisms of the numerous enzymesthat are currently grouped simply as phytase.Today, the term phytase is primarily based onthe in vitro capability of an enzyme to degrademyo-inositol hexakisphosphate (Konietzny andGreiner, 2002; see Greiner, Chapter 6, thisvolume). It does not identify the catalytic mecha-nism that is employed by the enzyme to hydrol-yse the substrate. Although there are a number ofphytases for which the mechanisms of myo-inositolhexakisphosphate hydrolysis are yet to be deter-

mined, the number for which this information isavailable is steadily increasing (Mullaney andUllah, 2003). It is thus logical to incorporate theresults of this research in a more informative clas-sification system. Differences in the physical fea-tures of phytases stem from molecular properties,and Oh et al. (2004) proposed a phytase classifica-tion system based on this fact. Physical character-istics of these enzymes, such as pH optima,mineral requirements, substrate requirementsand end product of hydrolysis, are all manifesta-tions of the distinct catalytic mechanisms.Different phosphatases have evolved to hydrolysemyo-inositol hexakisphosphate under diversifiedconditions.

Table 7.1 outlines a classification for phy-tases based on mechanistic enzymology. Thisclassification scheme continues the nomenclaturedeveloped by Oh et al. (2004), but incorporatesthe two classes of phytases, PAPhys and CPhys,that were not included in their system. Althougha large number of HAPs, PAPs and CPs cannotdegrade myo-inositol hexakisphosphate effectively,members of each class do share a common enzy-matic pathway with other members that hydrol-yse myo-inositol hexakisphosphate efficiently.Basing this system on enzyme family thereforeallows the incorporation of pertinent informationdeveloped by research on a significantly largernumber of enzymes. This classification systemalso offers the potential of assigning otheruncharacterized phytases based on unique fea-tures associated with individual enzyme families.The system is open-ended in that new groups ofphytases can easily be added and existing groupssubdivided when desirable.


Source Accession no. Consensus motif* * * * * * *

DXG GDXXY GNH(E/D) VXXH GHXH

A. niger JN0656 DMG GDLSY GNHE VLMH GHIH

G. max AF272346 DLG GDVTY GNHE VTWH GHVH

Fig. 7.3. Comparison of the amino acids composing the active-site consensus motif of Aspergillus nigerNRRL 3135 Apase and Glycine max GmPhy. Asterisks indicate the seven residues in the active sitecapable of forming ligands with metals. (From Schenk et al., 2000.)

106E

.J.Mullaney and A

.H.J.U

llah

Table 7.1. Classes of phytases.

Consensus motif or Organisms in unique structural which enzyme NCBI 3-D

Class Abbreviation Enzyme family feature occurs Unique characteristics model number

Histidine acid HAPhy Histidine acid RHGXRXP.. .HD Fungi, plants, Acid phosphatase; EDTA 1IHP phytase phosphatase bacteria stimulation; final product IP1 1QFX 1DKP

β-Propeller phytase BPPhy New family Six-bladed β- Bacteria (Plant Neutral to alkaline phosphatase; 1H6Lpropeller structure alkaline calcium required; final product IP3

phytases?)Cysteine phytase CPhy Cysteine phosphatase HCXXGXXR(T/S) Bacteria Acid phosphatase; inhibited by 1U24

Fe2+, Cu2+, Zn2+ and Hg2+;stimulated by Pb2+; final product inositol 2-monophosphate

Purple acid phytase PAPhy Purple acid DXG/GDXXY/GNH Plants Metalloenzymes None currently phosphatase (E,D)/VXXH/GHXH deposited

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Note

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8 Seed Phosphorus and theDevelopment of Low-Phytate Crops

Victor RaboyUnited States Department of Agriculture–Agricultural Research Service, Small Grains and Potato Germplasm Research Unit, 1691 S. 2700 W.,

Aberdeen, ID 83210, USA


Phytate (salts of myo-inositol hexakisphosphate)represents between 60% and 80% of mature seedtotal phosphorus (Raboy, 1997). Normal or ‘wild-type’ seeds produced by the major cereal cropssuch as maize (Zea mays L.), wheat (Triticum aes-tivum L.), barley (Hordeum vulgare L.) or rice (Oryzasativa L.) typically contain between 3.0 and4.0 mg P/g dry wt (Fig. 8.1, left; Raboy, 1997;Lott et al., 2000). Seeds produced by the majorlegume soybean (Glycine max L [Merr.]) typicallyhave a higher total phosphorus concentrationbetween 6.0 and 8.0 mg P/g dry wt. In bothcases, between 65% and 75% (±10%) of the totalphosphorus is found as phytate (illustrated for thecereal crops in Fig. 8.1, left). Inorganic phosphatenormally represents about 5% (±3%) of theremaining phosphorus, while lower-order myo-inositol phosphates usually represent <10%.

Phytic acid, the free-acid form of myo-inositolhexakisphosphate, is ubiquitous in eukaryotes. Itsmetabolism is important to a number of processesand functions in the eukaryotic cell, ranging fromphosphorus and mineral storage in seeds to signaltransduction, vesicular trafficking, stress response,RNA transport, DNA metabolism and the regula-tion of development (reviewed in Raboy, 2003;Shears, 2004). During seed development phytatemostly accumulates as mixed ‘phytin’ salts of sev-eral mineral cations. These are primarily mixedpotassium and magnesium salts, probably reflect-ing the relative abundance of these elements inthe seed. However, calcium, manganese, zinc and

iron phytates are also found (Lott et al., 1995).Phytate is often deposited as discrete globularinclusions called globoids, located within proteinstorage vacuoles (PSVs). In cereal grains, starchyendosperm PSVs primarily contain storage pro-tein deposits, whereas phytins are localized in thealeurone and germ (embryo and scutellum) PSVs.In maize, >80% of seed phytate is localized in thegerm, with the remainder in the aleurone. Insmall grains such as wheat, barley and rice, theopposite occurs; ≥80% of seed phytate is localizedin the aleurone, with the remainder in the germ(O’Dell et al., 1972). In the soybean seed phytindeposits are dispersed throughout the germ andcotyledonary tissues.

The primary applied interest in seed phytateconcerns its role in the nutritional quality of feedsand foods prepared from seeds. In the case ofanimal feeds, the primary interest is in phytate asthe seed’s major reserve of phosphorus. Non-ruminant (also known as monogastric) animalssuch as poultry, swine and fish do not efficientlydigest and utilize phytate (Brinch-Pedersen et al.,2002), which has two negative outcomes. First,non-ruminant feed must be supplemented withmineral phosphate to provide the animal’s nutri-tional requirement for phosphorus. Second, theexcretion of phytate can be an environmentalissue, as phosphorus in animal manure can con-tribute to water pollution (Sharpley et al., 1994;see Leytem and Maguire, Chapter 10, thisvolume). Reducing the environmental impact of

livestock production remains a major problem forproducers and society (Environmental ProtectionAgency, 2002). One approach to problems asso-ciated with feed phytate is to supplement feedswith the enzyme phytase (Brinch-Pedersen et al.,2002; see Lei and Porres, Chapter 9, this vol-ume). As ruminants such as beef and dairy cattletypically digest most feed phytate, it is generallynot considered a major issue in this context.

In contrast to the relatively straightforwardissues concerning feed phytate and phosphorusmanagement in non-ruminant production, theirrole in human nutrition and health are far morecomplicated. In the context of human nutrition,consumption of seed-derived phytate can con-tribute to mineral deficiencies, including iron,zinc, magnesium and calcium deficiencies. This isof particular concern to populations that dependon grains and legumes as staple foods, and espe-cially for women of childbearing age, infants andchildren within such populations (Brown andSolomons, 1991; Hurrell, 2003). However,dietary phytate may have a positive role as ananti-oxidant and anti-cancer agent (Graf et al.,

1987; Shamsuddin et al., 1988; Vucenik andShamsuddin, 2003; Singh and Agarwal, 2005;Somasundar et al., 2005) or as an inhibitor ofrenal stone formation (Grases et al., 2000). Theseissues remain unresolved and numerous ongoingstudies continue to address them.

Two observations need to be made concern-ing the issue of dietary phytate in human nutri-tion and health. First, the negative impacts ofdietary phytate are considered most important inrelatively younger people in the developing worldwho rely on cereal grains and legumes as staplefoods. In contrast, the potentially positive rolesappear to be most important to the health of age-ing individuals in developed countries. Thus, theissue of dietary phytate in human nutrition andhealth must be considered on a case-by-casebasis. Second, in the debate concerning theimportance of dietary phytate there is a generallack of cross-communication between the fields ofhuman and animal nutrition. In particular, itoften appears that those interested in the issue ofdietary phytate in human nutrition and healthignore the issue of dietary phytate in livestockproduction and the results of the numerous stud-ies on the impact of phytate in animal feed. Thisis unfortunate, because cereal or legume cropimprovement or production is only rarely anissue of producing food for people. Crop speciesmay often be used both in animal feed andhuman food.

The investigation of low phytic acid (lpa) geno-types of major crops began in the early 1990swith the isolation of two maize mutants (Raboyet al., 2000; Fig. 8.1, centre and right). These canbe used to develop ‘low-phytate’ crops, whichrepresent a second approach to addressing thedietary and environmental problems associatedwith seed phytate. In the seeds of these genotypes(Table 8.1), phytate is reduced by between 30%and 90%, but total phosphorus is typically notaltered to a great extent (a first exception to thisgeneral rule, barley lpa1-1, is discussed below).Instead, reductions in seed phytate are largelymatched by increases in inorganic phosphate(Fig. 8.1, lpa1-1), or in some cases by increasesin lower-order myo-inositol phosphates, suchas myo-inositol tetrakisphosphate or myo-inositolpentakisphosphate. These compounds containfour or five phosphate esters per molecule, ascompared with the six phosphates of myo-inositolhexakisphosphate (Fig. 8.1, centre and right).

112 V. Raboy

0

See

d ph

osph

ate

frac

tion

(mg

P/g

)

2

Wild-type

Maize genotype

Inorganic P

Other P

Phytate

Lower inositol P

Ipa1-1 Ipa2-1

4

Fig. 8.1. Typical seed phosphorus fractions inthree types of maize genotypes: homozygous wild-type or ‘normal’; low phytic acid 1-1; low phytic acid2-1. ‘Lower inositol P’ refers to inositol phosphateswith five or fewer phosphate esters per moleculecompared with six in phytate. ‘Other P’ refers to allforms of phosphorus in seeds other than in phytate,lower inositol phosphates or inorganic phosphates,such as DNA, RNA, protein and phospholipids.

The biochemistry and molecular genetics of phy-tate metabolism has also continued to progressgreatly. Many of the enzymes involved in thephosphorylation of myo-inositol to the hexak-isphosphate have been described and genesencoding such enzymes have been identified. Insome cases, lpa genotypes have been used in ‘for-ward genetics’ approaches to identify genes andfunctions important to seed phytate synthesis(Shi et al., 2003, 2005a). Recent studies usingArabidopsis, the model organism for much plantscience research, are also contributing advancesto this field (Stevenson-Paulik et al., 2002, 2005).

In contrast, relatively little progress has beenmade in the molecular biology of seed total phos-phorus, defined as the sum of all forms ofphosphorus in the seed. Manipulating theamount of phytate in seeds, such as in lpa geno-types, has not greatly influenced seed total phos-phorus. In lpa genotypes the chemistry of thephosphorus is altered, but not the total concen-tration. Reduced total phosphorus may havevalue when seed crops are used in ruminant(dairy and beef) feed (Volk et al., 2000; Erickson

et al., 2002; Rotz et al., 2002; Toor et al., 2005; seeDao, Chapter 11, this volume), as well as in otherapplications, including non-ruminant feed. Whilea great deal of progress has been made in under-standing the molecular biology of the uptake anddistribution of phosphorus in the parent plant,less is known about the uptake and distribution ofphosphorus in the seed.

This chapter reviews the biochemistry andgenetics of seed phytate and lpa genotypes andthe breeding and nutritional evaluation of ‘low-phytate’ crops. It concludes with a look at futuredirections for research on seed phosphate.

Metabolic Pathways, Genesand Mutants

Substantial progress has been made in the molecu-lar genetics and biochemistry of phytic acid biosyn-thesis, which represents one component of themyo-inositol phosphate pathways. In developingseeds the myo-inositol phosphate metabolic

Seed Phosphorus and Low-Phytate Crops 113

Table 8.1. Seed phosphorus fractions in maize, barley, rice, wheat and soybean low phytic acidgenotypes.a

Species Genotype Total P (g P/kg) Total inositol P Inorganic P Cellular Pb

Maize Wild-type 4.5 3.4 0.3 0.8Maize lpa1-1 4.7 1.1 3.1 0.5Maize lpa2-1 4.6 2.6 1.3 0.7Barley Wild-type 4.8 2.9 0.4 1.5Barley lpa1-1 (M 422) 3.7 1.2 1.2 1.3Barley lpa2-1 (M 1070) 5.4 1.9 1.7 1.7Barley lpa 3-1 (M 635) 5.0 0.7 2.5 1.8Barley M 955 5.0 NDc 3.3 1.7Rice Wild-type 3.1 2.23 0.14 0.79Rice lpa1-1 3.5 1.37 1.13 1.05Wheat Wild-type 5.3 4.0 0.5 0.8Wheat JS-12-LPA 5.1 2.5 2.6 NDc

Soybean Wild-type 7.95 5.68 0.30 1.97Soybean M 153 7.96 1.98 3.02 2.96

aAll analyses listed in this table were conducted in the USDA–ARS Cereal Chemistry (Raboy) laboratory. (From Larsonet al., 2000; Raboy et al., 2000; Wilcox et al., 2000; Dorsch et al., 2003; Oltmans et al., 2005 using methods asdescribed.) Total inositol phosphate is primarily myo-inositol hexakisphosphate, but the analytical method used will alsodetect lower-order inositol phosphates if they are present. Cellular phosphorus includes all forms of phosphorus otherthan inositol phosphate and inorganic phosphate, including phosphorus in starch, phospholipids, RNA and DNA.bCellular phosphorus = total phosphorus – (inositol phosphate + inorganic phosphate).cND = not detected. Inositol phosphate concentrations in M 955 were below that which is reliably assayed with themethods used. In the case of the wheat JS-12-LPA mutant, in the analyses given in this table, the sum of inositolphosphate and inorganic phosphate equalled total phosphorus, indicating no other forms of cellular phosphorus. This isclearly an artefact of this particular assay.

pathways can be viewed as consisting of two parts:(i) synthesis and/or supply of myo-inositol and phos-phate (Fig. 8.2, top); and (ii) myo-inositol phos-phate/phosphatidylinositol (PtdIns) phosphatemetabolism leading to myo-inositol trisphosphatesand ultimately phytic acid (Fig. 8.2, middle andbottom). The enzyme D-myo-inositol 3-monophos-phate synthase (MIPS) is the sole synthetic source

of the myo-inositol ring (Loewus and Murthy,2000). Its activity is coupled with myo-inositol3-monophosphatase (IMP) in a simple MIPS/IMP‘myo-inositol synthesis pathway’. Hitz et al. (2002)demonstrated that the LR33 mutation in a soybeanMIPS gene resulted in a block in seed phytateaccumulation, indicating that a substantial fractionof the myo-inositol necessary for phytate synthesis in

114 V. Raboy

Phosphatidylinositol

phosphate earlyintermediate

pathway

Inositolphosphate

earlyintermediate

pathway

Ins(3,4,6)P3

PtdIns(4,5)P2

Ins(1,4,5)P3Ins(1,3,4)P3

PtdIns(4)P15-kinase

Phospholipase C

H

H

HH

H

OH

OH

HO1

234

6P

PH

P

5P

H

H

H

H

H

OH

HO1

234

6

PH

P

5

PP

H

H

H

H

H

1

234

6

PP

H

P

5

Ins(1,2,3,4,5,6)P6(phytic acid)

P

P

H

H

H

H

H

OH

OH

OH

HO

HO

H

1

234

5 6

H

H

H

H

H OH

OH

OHH

1

234

5 6P

PP

H

H

H

H

H

OH

OH

OH

OH

HO

HO

H

1

234

5 6

More highly phosphorylatedIns hepta- and octa-phosphatesInsP7 or InsP8

Breakdown viaphytases

Inositol polyphosphatekinases

Glucose 6-phosphate

Ins(3)P1 synthase

Inositolmonophosphatase

Inositolkinase

PtdIns(4)P1

PtdIns4-kinase

PtdInsmyo-Inositol (Ins)Ins(3)P1

Phosphatidylinositolsynthase

(PtdIns synthase)

PP

H

H

H

H

H

1

234

6

P

H

P

5P

OH

Ins(1,3,4,5,6)P5

Inositol polyphosphate2-kinase

Ins(1,3,4,5)P4

Inositol polyphosphatekinases

Soybean LR33 mutation(Hitz et al., 2002)

Maize lpa2(Shi et al., 2003)

Maize lpa3(Shi et al., 2005b)

Arabidopsis IPK2 mutants(Stevenson-Paulik et al., 2005)

Arabidopsis IPK1 mutants(Stevenson-Paulik et al., 2005)

Fig. 8.2. Biochemical pathways leading from glucose 6-phosphate to myo-inositol (Ins) and ultimately tomyo-inositol hexakisphosphate. Steps in the pathways for which there are known mutations or blocks areindicated by crossing lines accompanied by the published reference describing the mutation(s). The sixcarbons of the myo-inositol ring are numbered according to the D-numbering convention. P represents aphosphate group, PH2O4.

seeds is synthesized de novo. Yoshida et al. (1999)had previously demonstrated that MIPS isexpressed proximal to the site of phytate accumula-tion during the development of rice seeds.

Pathways to phytic acid might then eitherproceed through sequential phosphorylation ofmyo-inositol and ‘soluble myo-inositol phosphates’(Fig. 8.2, middle-left) or by one of two alternativepathways that utilize PtdIns early intermediates(Fig. 8.2, middle-right). In the pathway proposedby Biswas et al. (1978a), myo-inositol 3-monophos-phate is directly converted to myo-inositol1,3,4,5,6-pentakisphosphate through sequentialphosphorylation by a phosphoinositol kinase, ofwhich two electrophoretic forms were identified.The conversion of myo-inositol 1,3,4,5,6-pentak-isphosphate to phytic acid is then catalysed by amyo-inositol hexakisphosphate–adenosine diphos-phate phosphotransferase (Biswas et al., 1978b),now commonly referred to as myo-inositol1,3,4,5,6-pentakisphosphate 2-kinase (Phillippyet al., 1994). An alternative pathway to phyticacid begins with myo-inositol as the initial sub-strate and myo-inositol kinase activity (E.C.2.7.1.6.4; English et al., 1966; Loewus et al., 1982)and proceeds through site-specific sequentialphosphorylation steps of defined soluble myo-inositol phosphates (Fig. 8.2, middle-left). Thispathway was described in studies of the cellularslime mould Dictyostelium discoideum (Stephens andIrvine, 1990) and the monocot Spirodela polyrhiza(Brearley and Hanke, 1996b). These ‘inositolphosphate early intermediate’ (Fig. 8.2, left side)or ‘lipid-independent’ (Stevenson-Paulik et al.,2002) pathways are similar, and share the com-mon intermediate myo-inositol 3,4,6-trisphosphate(Fig. 8.2). Recently, Shi et al. (2005) determinedthat the maize lpa3 gene encodes myo-inositolkinase, providing the first genetic evidence for theexistence of myo-inositol kinase and demonstrat-ing its importance to phytate synthesis and accu-mulation in seeds.

The pathways to phytic acid that involvePtdIns phosphate lipid intermediates and myo-inositol 1,4,5-trisphosphate are illustrated inFig. 8.2 (middle-right). In a pathway to phyticacid expressed in the nucleus of yeast (York et al.,1999), PtdIns 4,5-bisphosphate is hydrolysed toyield myo-inositol 1,4,5-trisphosphate, which isthen phosphorylated directly to myo-inositol1,3,4,5,6-pentakisphosphate by myo-inositol 1,4,5-trisphosphate 3/6-kinase that is encoded by the

IPK2 gene. A variety of names have been usedfor this and other types of inositol phosphatekinases, but for simplicity it is referred to hereonly as myo-inositol 1,4,5-trisphosphate 3/6-kinase. Similarly, the second type of multifunc-tional myo-inositol polyphosphate kinase discussedimmediately below is referred to only as myo-inos-itol 1,3,4-trisphosphate 5/6-kinase. Stevenson-Paulik et al. (2002) isolated two Arabidopsis thalianagenes related to the yeast myo-inositol 1,4,5-trisphosphate 3/6-kinase, termed AtIpk2α andAtIpk2β. Stevenson-Paulik et al. (2005) recentlydemonstrated that mutations in AtIpk2β impacton the ability of seeds to synthesize phytic acid,indicating that the pathway does proceed at leastin part via the intermediate myo-inositol 1,4,5-trisphosphate.

Alternatively, studies of human cells describeda pathway whereby myo-inositol 1,4,5-trisphosphateis first converted to myo-inositol 1,3,4,5-tetrakispho-sphate and then to myo-inositol 1,3,4-trisphosphate(Fig. 8.2, centre), which is subsequently phosphory-lated to myo-inositol 1,3,4,5,6-pentakisphosphatethrough what was first defined as myo-inositol 1,3,4-trisphosphate 5/6-kinase (Wilson and Majerus,1996, 1997). The maize lpa2 gene encodes myo-inositol 1,3,4-trisphosphate 5/6-kinase (Shi et al.,2003). In seeds produced by plants homozygous formaize lpa2 null mutations, phytic acid is reducedby 35%, clearly indicating that this second type ofmyo-inositol polyphosphate kinase is important toseed phytic acid synthesis. myo-Inositol 1,3,4-trisphosphate and myo-inositol 1,4,5-trisphosphatekinases, such as those encoded by maize lpa2 andArabidopsis AtIpk2β, respectively, can phosphorylatemultiple myo-inositol phosphates (reviewed inShears, 2004). One or perhaps both types of myo-inositol polyphosphate kinases working togethermight be able to convert myo-inositol 3,4,6-trisphos-phate to myo-inositol 1,3,4,5,6-pentakisphosphate.Thus, a pathway to phytate that proceeds entirelyvia soluble myo-inositol phosphates might in factutilize enzymes considered part of the PtdIns-inter-mediate pathway to phytate (Stevenson-Pauliket al., 2002; Raboy, 2003).

In summary, it is of interest that the threekinase mutations that perturb phytic acid accu-mulation in seed (maize lpa3, maize lpa2 andArabidopsis Ipk2) represent genes encoding func-tions that represent markers for the three alterna-tive pathways to phytate. Maize lpa3 mutantsblock myo-inositol kinase activity believed to catal-


yse the first step in the soluble myo-inositol phos-phate/lipid-independent pathway; ArabidopsisIPK2 mutants block myo-inositol 1,4,5-trisphos-phate 3/6-kinase activity, critical to the type oflipid-dependent pathway that proceeds directlyfrom myo-inositol 1,4,5-trisphosphate to myo-inosi-tol 1,3,4,5,6-pentakisphosphate; maize lpa2mutants block myo-inositol 1,3,4-trisphosphate5/6-kinase activity, believed to be critical toan alternative lipid-dependent pathway firstdescribed in the studies of human cells. This indi-cates that either the pathway to phytic acid inseeds is complex and non-linear or that ourunderstanding and definition of these genes andenzymes requires revision. Regardless of precur-sor pathways, there is a consensus that myo-inosi-tol 1,3,4,5,6-pentakisphosphate is the penultimatemyo-inositol phosphate in the pathway to phyticacid, and that its conversion is catalysed by myo-inositol polyphosphate 2-kinase (Biswas et al.,1978a,b; York et al., 1999). Stevenson-Paulik et al.(2005) demonstrated that a mutation in anArabidopsis gene encoding a 2-kinase substantiallyblocked seed phytic acid accumulation.

The existence of myo-inositol phosphatesmore highly phosphorylated than phytic acidseems to occur relatively widely in eukaryoticcells (Stephens et al., 1993; Laussmann et al.,2000). These compounds contain pyrophos-phate moieties and include 5-diphosphoinositol

1,2,3,4,6-pentakisphosphate (with seven phos-phate groups) and bis-diphosphoinositol 1,2,3,4-tetrakisphosphate (with eight phosphate groups).To date there has been very little progress in thestudy of these pyrophosphate-containing com-pounds in plant systems (Brearley and Hanke,1996a; Flores and Smart, 2000; Dorsch et al.,2003). Their potential role in seed phytatemetabolism, localization or deposition isunknown.

Phytate Deposition in Globoids

While this brief review indicates that muchprogress has been made in the molecular biologyof the structural synthetic pathway leading tophytic acid synthesis, relatively little progress hasbeen made regarding how phytate salts areformed and deposited as globoids. Little is knownabout how the proteins, substrates and productsimportant to these pathways and processes arelocalized and compartmentalized within the cell.Figure 8.3 provides a model for a phytate-accumulating PSV or membrane-bound globoid,illustrating several of the localization and trans-port functions of possible importance. An ongoing debate concerns whether or not individ-ual globoids are membrane-bound. Jiang et al.

116 V. Raboy

Fig. 8.3. Components of a membrane-bound organelle, representing either a protein storage vacuole(PSV) or a membrane-bound globoid found within a compound PSV. V-PPase = vacuolar inorganicpyrophosphatase; V-ATPase = vacuolar adenosine triphosphatase; TIP = tonoplast intrinsic protein;PtdIns = phosphatidylinositol. Question marks indicate speculative aspects of the diagram.

(2001) provided evidence that globoids are mem-brane-bound, and therefore that the PSVs thatcontain them represent a ‘compound organelle’.For the purpose of this discussion, Fig. 8.3 illus-trates an organelle defined by a single mem-brane. The PtdIns phosphates that might serve asintermediates in the synthesis of phytic acid arethemselves localized to the membrane that bindsthe phytate-accumulating PSV.

Specific molecular details of how mixed phy-tate salts are deposited within seed PSVs areunknown. Water is probably transported into thePSV by aquaporins, one of the several isoforms oftonoplast intrinsic proteins. One or more suchproteins might be specific to the globoid contain-ing PSV ( Jauh et al., 1999; Takahashi et al., 2004).Second, the PSV (or globoid) membrane probablycontains both vacuolar pyrophosphatases (V-PPase) and adenosine triphosphatase, which breakdown inorganic pyrophosphate or adenosinetriphosphate, respectively, and pump protons intothe internal vacuolar space (Maeshima, 2000;Jiang et al., 2001). This establishes a proton gradi-ent that drives transport, through channels andantiporters, of various solutes from the cytoplasminto the PSV. These solutes must include potas-sium and magnesium counter-ions, and possiblyother minerals. Adenosine triphosphate–bindingcassette transporters (ABCs), which use the energyprovided by adenosine triphosphate breakdown totransport a variety of solutes ( Jasinski et al., 2003),play some role since maize lpa1 encodes an ABCtransporter (Shi et al., 2005a).

As transport functions are probably import-ant to seed phytate deposition, mutations ingenes encoding such functions probably wouldimpact net seed phytate accumulation. The firstexample of a single-gene mutation is a transportfunction that impacts seed phytate is Maize lpa1(Shi et al., 2005a). Quantitative variation in thelevels of phosphate and phytate in vegetative andseed tissues of Arabidopsis was used to identify aquantitative trait locus (QTL) that accounts for asignificant amount of the variation observed(Bentsink et al., 2003). Contained within the 99-kb chromosomal segment represented by thisQTL were 13 open reading frames, one of whichencoded a putative vacuolar adenosine triphos-phatase (V-ATPase). Bentsink et al. (2003)hypothesized that the variation in phosphate andphytate levels observed among the Arabidopsislines in their study was in large part due to varia-

tion in phosphate transport caused by heritabledifferences in this enzyme.

Low-Phytate Crops

The first two lpa mutations were isolated in maize(maize lpa1-1 and maize lpa2-1; Fig. 8.1; Raboyet al., 2000) by screening seed sampled fromchemically mutagenized populations directly forreduced seed phytic acid, using a paper elec-trophoresis method. Analyses of seed producedby plants homozygous for either maize lpa1-1 orlpa2-1 indicated that the reductions in phytic acidwere largely matched by increases in inorganicphosphate, so that total phosphorus in the seedremained unchanged and similar to the wild type(Fig. 8.1; Table 8.1). The increase in seed inor-ganic phosphate in these first two lpa mutants wasseveral-fold, from less than 0.5 mg P/g in non-mutant seed to between 1.0 and 3.0 mg P/g inthe mutants. As accurate tests for seed inorganicphosphate are much quicker and straightforwardthan methods to assay phytic acid (such as thoserequiring precipitation, purification or chro-matography), and as the increase in seed phos-phate in lpa mutants compared with the wild typeranges up to tenfold, a simple high-throughputscreen that tests for the ‘high inorganic P’ seedphenotype of lpa genotypes can be readily designedfor use in genetics studies. This assay has subse-quently been used to isolate mutants that defineda third lpa locus in maize (Shi et al., 2005b), andto isolate lpa mutations in barley (Larson et al.,1998; Rasmussen and Hatzack, 1998), rice(Larson et al., 2000), soybean (Wilcox et al., 2000)and wheat (Guttieri et al., 2004). This same high-throughput high inorganic P test also facilitatesgenetic mapping, and has been used to map lpaloci in maize (Raboy et al., 2000), barley (Larsonet al., 1998) and rice (Larson et al., 2000; Andayaand Tai, 2005).

lpa Mutations have been used to developfirst-generation germplasm useful in initial evalu-ations of the agronomic properties and nutri-tional value of low-phytate types. The approachtaken in these first studies was to use lpa muta-tions and standard ‘backcrossing’ breeding meth-ods to develop sets of ‘near-isogenic’ lines. Thesesets consist of sibling lines that are eitherhomozygous for a given wild-type non-mutant


allele, and produce seed with normal levels ofphytate and inorganic phosphate, or homozygousfor the given lpa allele and produce seed withreduced phytate and increased inorganic phos-phate. These sibling lines are referred to as ‘near-isogenic’ because their genomes are similar, otherthan allelic differences at an lpa gene. In theory,standard backcrossing methods involving a mini-mum of six crosses should mean that identicalalleles are shared at >98% of all genes. Therefore,any differences observed in plant or seed pheno-type and agronomic performance, or differencesin nutritional quality, can in theory be attributedlargely to the allelic difference at a single lpa gene.

These types of near-isogenic lines were firstproduced in maize using the lpa1-1 allele of themaize lpa1 gene. Plants homozygous for lpa1-1produce seed with a phytate reduction of about66% compared with seed produced by wild-typenon-mutant sibling lines (Table 8.1). As hybridseed is used for maize production, near-isogenicsets of inbred lines were first produced, whichwere then used to produce 14 different pairs ofnear-isogenic hybrids (isohybrids; Ertl et al.,1998). Field trials of these first isohybrid pairsindicated that yields were reduced in eight of thelpa1-1 hybrids compared with their matchingwild-type isohybrids, but yields of the lpa1-1 iso-hybrid in the remaining six pairs were similar tothe wild type. Overall a yield reduction of about6% was observed in the lpa1-1 hybrids, although

major differences in other aspects of plant growthand seed function were not observed.

Pairs of sibling near-isogenic lines weredeveloped using four barley lpa mutations: lpa1-1(formerly known as M 422), lpa2-1 (formerly M1070), lpa3-1 (formerly M 635) and M 955.Homozygosity for these mutations results in seedphytate reductions ranging from moderate(35–50%, lpa1-1 and lpa2-1) to relatively large(70–80%, lpa3-1) and extreme (>90%, M 955)(Table 8.1). Yield of isolines representing wild-type and mutant siblings were evaluated in fieldlocations in Idaho, USA, during 2 years ofregional drought (Bregitzer and Raboy, 2006;Table 8.2). These production environments wereeither irrigated, which resulted in relatively lowdrought stress, or non-irrigated (‘dryland’ or‘rain-fed’), which resulted in considerabledrought stress. The yield of isolines representinglpa1-1, lpa2-1 and lpa3-1 were statistically similarto wild-type controls when grown with irrigation.However, the yield of isolines representing the‘extreme reduction’ mutant M 955 was clearlyreduced, even in the relatively stress-free, irri-gated production environments. Yield reductionsassociated with lpa mutations were more pro-nounced in the more stressful non-irrigated envi-ronments. In these environments only the yield ofbarley lpa1-1 was statistically indistinguishablefrom its wild-type sibling lines. The yield of iso-lines representing lpa2-1, lpa3-1 and M 955 were

118 V. Raboy

Table 8.2. Comparison of yield of barley wild-type and low phytic acid sibling isolines when grown innon-stressful (irrigated) and stressful (dryland) environments.a

Seed phytic acid Yield (kg/ha)c

Sibling pair Genotypeb reduction vs. wild-type(%) Irrigated Dryland

1 Wild-type − 8320 A 1935 A1 lpa1-1 50 % 8487 A 1718 A2 Wild-type − 8429 A 1728 A2 lpa2-1 33 % 8271 A 1321 B3 Wild-type − 7994 A 1836 A3 lpa3-1 70 % 7991 A 1353 B4 Wild-type − 8253 A 1769 A4 M 955 90 % 7147 B 1162 B

aIrrigated environments were fields at Aberdeen, Idaho, in 2002 and 2003, and Filer, Idaho, in 2003. Drylandenvironments were fields at Tetonia, Idaho, in 2002 and 2003, and Soda Springs, Idaho in 2003. Two replicates of sixlines representing each genotype were grown at each location in a randomized complete block.bAll barley lines were in the cultivar Harrington genetic background.cValues followed by the same letter within each sibling pair are not significantly different (P = 0.05).

all substantially reduced, compared with wild-type sibling lines, when grown under droughtstress. Importantly, the reductions appeared pro-portional to the reduction in seed phytate.

These results clearly indicate that lpa muta-tions can be associated with yield losses anddecreased stress tolerance, but that such lossesare variable and are probably both function-and/or gene/allele-specific. That is, perhapsmutations in different types of functions mighthave a much smaller impact on yield or stresstolerance than those in other functions, genes oralleles of a given gene. Such mutations couldinclude those in myo-inositol phosphate synthesisvs. mutations in substrate or phosphorus trans-port, or those giving rise to different alleles of agiven gene.

Barley lpa1-1 represents one case in whichcarefully conducted and replicated field trials indi-cate yields comparable to non-mutant siblinglines, possibly even in relatively stressful environ-ments. Interestingly, the barley lpa1-1 mutationaffects phytate accumulation only in the barleyaleurone layer – the outer layer of the cereal grainendosperm (Ockenden et al., 2004). In cerealgrains phytate accumulates in germ tissue (includ-ing the embryo and scutellum) and the aleurone,but not in the central endosperm. In seedshomozygous for barley lpa1-1, only aleurone phy-tate is reduced; germ phytic acid is similar tothe wild type. In contrast, in seeds homozygousfor barley lpa2-1, lpa3-1 and M 955, both germand aleurone phytate are similarly reduced.Therefore, the relatively good field performanceof barley lpa1-1 might be due to the fact that thismutation is in a gene of limited tissue specificity,thus reducing the impact on plant growth andperformance. Alternatively, lpa1-1 might be amutation in a transport function as opposed toa mutation in the myo-inositol phosphate pathways,and thus have a more limited impact on functionsimportant to signal transduction and stressresponse, which in turn are important to yield.

lpa Mutations directly impact on three cellu-lar pools important to numerous pathways: phos-phorus, myo-inositol and myo-inositol phosphates.Greatly altering these important metabolic poolscould lead to yield loss. The reduced yields of lpavariants of maize and barley, compared withwild-type controls, could be due in part to a nega-tive impact of high levels of seed inorganic phos-phate on starch synthesis and accumulation.

Starch represents the major component (>50%)of the dry weight in cereal grains, so any factorimpacting on seed starch accumulation would alsoimpact on yield. For example, a rate-limiting stepin seed starch synthesis is catalysed by the enzymeadenosine diphosphate–glucose pyrophosphory-lase (AGP; E.C. 2.7.7.27), which is allostericallyinhibited by a high inorganic phosphate concen-tration (Hannah, 1997). A genetically engineeredversion of this enzyme that is insensitive to inor-ganic phosphate increased seed yield and plantbiomass in rice (Smidansky et al., 2003). A usefulapproach to developing high-yielding lpa cultivarsor hybrids might therefore be to engineer thesetypes to express ‘deregulated’ AGP in seeds.

In addition to metabolic impacts, thesemutations probably have numerous downstreameffects on functional properties such as stress tol-erance (discussed above). Simply isolating muta-tions or alleles that only impart the low-phytatetrait represents a first step. In addition to possibleapproaches that involve genetic engineering,developing high-yielding, stress-tolerant low-phytate crops will probably require classicalbreeding techniques, such as recurrent selectionfor yield and performance within lines that arehomozygous lpa. This would select for combina-tions of ‘favourable’ alleles at numerous loci thatmodify or reduce the impact of the lpa genotype.

This process has begun with soybeans.A soybean low-phytate mutant termed M 153 wasisolated (Wilcox et al., 2000; Table 8.1) and subse-quently shown to require the inheritance of reces-sive alleles at two non-linked loci, termed pha1and pha2 (Oltmans et al., 2004). Crosses to severalsoybean lines resulted in the development of threepopulations (Oltmans et al., 2005). Ten wild-typelines (homozygous for dominant wild-type alleles)and ten low-phytate lines (homozygous for bothpha1 and pha2 alleles) were isolated within eachpopulation. Analysis of seed traits and field per-formance found relatively little statistically signifi-cant differences between normal and low-phytatelines, except that the latter displayed reduced fieldemergence compared with sibling wild-type lines(45% vs. 68%). However, variation in field emer-gence within low-phytate lines indicates that posi-tive selection may yield low-phytate soybeangermplasm with acceptable field emergence.

In many cases analyses of seed phosphorusfractions in lpa genotypes indicate that reductionsin phytate are largely matched by increases in


inorganic phosphate or by increases in both inor-ganic phosphate and other lower-order myo-inosi-tol phosphates. This means that cellularphosphorus, defined as all forms of phosphorusother than myo-inositol phosphates and inorganicphosphate, tends to remain constant (Table 8.1).However, in some cases, such as the soybeanmutant M 153 (Table 8.1), the reduction in phy-tate phosphorus is not matched entirely byan increase in inorganic phosphate and othermyo-inositol phosphates (Oltmans et al., 2005),meaning that cellular phosphorus is increasedcompared with the wild type. Further analysesare required to identify the phosphorus com-pound(s) involved.

A second study (Meis et al., 2003) revealedan intriguing downstream effect of a second typeof low-phytate soybean. Hitz et al. (2002) isolateda mutant, LR33, subsequently shown to be arecessive mips allele in an MIPS loci in the soy-bean genome. Meis et al. (2003) observed areduction in seedling emergence for low-phytatelines homozygous for the LR33 mips allele.However, reduced field emergence of LR33 linescompared with the wild type was influenced byseed production environment: LR33 seed pro-duced in temperate locations such as Iowa, USA,had 63% field emergence, while seed from thesame lines produced in subtropical environmentssuch as Puerto Rico had a mean field emergenceof 8%. In comparison, seed produced by wild-type sibling lines in temperate locations had amean field emergence of 77% compared with83% for seed produced in the subtropical envi-ronment. Homo-zygosity for the LR33 recessivemips allele therefore has an interesting and agro-nomically important downstream effect, becauseproduction of LR33 seed in subtropical environ-ments results in seed with greatly reduced fieldemergence, even when germinated and grown intemperate locations. The cause of this effect is atpresent unknown.

Animal Nutrition Studies EvaluatingLow-Phytate Crops

In terms of animal nutrition and production, themain interest in lpa crops focuses on grain phos-phorus availability in diets for non-ruminantspecies such as poultry, swine and fish. The

primary concerns are providing sufficient phos-phorus for optimal animal growth and productiv-ity, while reducing phosphorus concentrations inanimal manure. Also of interest are calciumavailability and several other measures of animalhealth, productivity and product quality. Perhapsthe best measure of phosphorus availability infeed is the difference between phosphorus intakein feed and phosphorus excretion in manure.Phosphorus availability is also often measuredindirectly in animals fed diets prepared with nor-mal vs. low-phytate grains, by comparing variousmeasures of animal growth rate, feed/gain ratios,bone ash and strength, and concentrations ofphosphorus or calcium in bones or blood.

The first published study (Ertl et al., 1998)addressing these questions compared the nutri-tional value of grain produced by two maize iso-hybrids in diets for broiler chicks. This involvedgrain from a ‘normal phytate’ wild-type isohybridcontaining 3.8 mg total P/g dry weight (3.2 mgP/g as phytate) and a matching lpa1-1 isohybridcontaining 3.9 mg total P/g dry weight (1.3 mgP/g as phytate). If we hypothesize that phytate islargely ‘non-available phosphorus’ to non-ruminants, and all other forms of seed phosphorus(referred to as non-phytate phosphorus) representavailable phosphorus, then the wild-type graincontained about 0.6 mg P/g as available phos-phorus or 16% of grain total phosphorus, and thelpa1-1 grain contained 2.6 mg P/g as availablephosphorus or 67% of grain total phosphorus.

The maize wild-type and lpa1-1 experimentaldiets utilized in the Ertl et al. (1998) chick feedingtrial were formulated identically, so the only dif-ference between the two diets was in their phos-phorus chemistry. Differences in responses todiets were therefore attributable to differentialavailability of phosphorus. Two diets wereprepared for each of the two genotypes, whereinmaize grain represented either 56% or 69% ofthe total diet. The additional, ‘basal’ componentsof the diets (soymeal, vitamins, minerals, anti-biotics, etc.) contributed to about 25% of thetotal phosphorus, and no additional supplemen-tal phosphorus was included. The maize compo-nent therefore represented the major source ofphosphorus in the diets. All diets were otherwiseformulated to contain similar, National ResearchCouncil-recommended levels (<5% difference) ofother, non-phosphorus constituents providingsimilar energy, essential amino acids and

120 V. Raboy

minerals, including calcium (National ResearchCouncil, 1994).

Estimates of available phosphorus in grainproduced by the wild-type isohybrid rangedbetween 30% and 48%, or about 1.1–1.8 mg P/g(Ertl et al., 1998). The corresponding estimates ofavailable phosphorus for the maize lpa1-1 grainwere between 70% and 91%, or about 2.7–3.5mg P/g. These differences parallel those in non-phytate phosphorus obtained by chemical analy-ses of the grains, confirming that non-phytatephosphorus is largely available to non-ruminants.As the total phosphorus concentration in grain isoften similar in normal and low-phytate varieties,and phytate phosphorus normally represents asubstantial fraction of grain total phosphorus,low-phytate genotypes have greatly increasedavailable phosphorus.

Optimal chick growth under the experimen-tal conditions used in the Ertl et al. (1998) studywas obtained with experimental diets preparedusing monosodium phosphate as the phosphorussource to provide 0.4–0.5 mg P/g dry feed, reflect-ing current recommendations (National ResearchCouncil, 1994). Therefore, the diets prepared withwild-type maize provided only 20–40% of thephosphorus required for optimal growth, and thislimited bird growth (bird weights at 18 days were64–71% of those obtained from diets containing100% recommended phosphorus). The diets for-mulated with lpa1-1 grain provided between 50%and 70% of the optimal level of available phos-phorus, still limiting to optimal growth but less sothan wild-type grain (bird weights at 18 days were76–84% of those obtained from diets containing100% recommended phosphorus).

The objective in engineering optimizednutrient chemistry for animal agricultural pro-duction is to maximize performance or produc-tion while minimizing waste. Faecal phosphorusfrom birds consuming lpa1-1 maize diets wasreduced by between 9% and 40% compared withthat from birds consuming wild-type maize diets,and by between 30% and 47% compared withthat from birds fed diets containing phosphatesupplements (Ertl et al., 1998).

Phosphorus nutritional status has an impor-tant impact on calcium status and bone health. Itis significant that blood phosphorus and calciumlevels in birds consuming the lpa1-1 maize dietswere 46% and 49% greater, respectively, than inbirds consuming the wild-type maize diets (Ertl

et al., 1998). This resulted in increases of 12%,10% and 13% in bone (tibia) ash, bone phospho-rus and bone calcium, respectively. The observeddifferences in the availability of phosphorus andcalcium and the health and productivity of theanimals can be attributed to the major differ-ences in grain phosphorus chemistry between thewild-type and lpa1-1 grain, in turn attributable inlarge part to a single allelic difference in the twomaize genotypes.

This first animal feeding trial of a low-phy-tate maize grain was followed by a number ofstudies that either confirmed or expanded uponthe initial results described above: available phos-phorus is increased in low-phytate grains inproportion to the increase in non-phytate phos-phorus; depending on dietary formulation, use oflow-phytate grains reduces animal waste phos-phorus in proportion to reduced dietary phytate,with faecal phosphorus reductions between 10%and 50%; and reduced dietary phytate canenhance availability of calcium, iron and zinc.These studies used a variety of experimentalapproaches, including diets formulated witheither low-phytate maize or low-phytate barleys,diets that evaluated products made from thesegrains such as low-phytate maize gluten, dietsformulated using combinations of low-phytategrains and phytase enzyme supplements, and sev-eral animal systems including fish (trout), swineand poultry (chicken and turkey) (Sugiura et al.,1999; Douglas et al., 2000; Li et al., 2000, 2001;Spencer et al., 2000b; Waldroup et al., 2000;Veum et al., 2001, 2002; Peter and Baker, 2002;Jang et al., 2003; Overturf et al., 2003; Thackeret al., 2003; Yan et al., 2003).

A recent development in this field was theevaluation (Sands et al., 2003; Adeola, 2005; Karr-Lilienthal et al., 2005) of the nutritional value ofsoymeal made from the M 153 low-phytate soy-bean described in Wilcox et al. (2000). Most of thephosphorus and phytate in the seed are concen-trated in soymeal when it is processed from wholesoybeans (V. Raboy, 2006, unpublished data).Phosphorus availability is increased in low-phytatesoymeal compared with normal phytate soymeal,in proportion to the increase in non-phytate phos-phorus (Sands et al., 2003). These studies alsofound that ‘metabolizable energy’ and amino aciddigestibility are also increased in low-phytatesoymeal compared with normal soymeal (Adeola,2005; Karr-Lilienthal et al., 2005).


The first published trout feeding trial(Sugiura et al., 1999) was also the first to evaluate,side by side in the same study, diets formulatedwith low-phytate maize (two lpa1-1 isohybridswith phytate reductions of about 66%) or low-phytate barley (an isoline of the cultivarHarrington homozygous for the barley lpa1-1allele, formerly referred to as M 422, whichresults in about a 40% reduction in grain phytate;Table 8.1). This study demonstrated that whendiets were formulated to contain relatively high orluxury levels of available phosphorus, most fromnon-grain components such as fishmeal, little ben-efit in terms of phosphorus or mineral availability,or reduction in waste phosphorus, was observedwith the substitution of low-phytate grains fornormal grains. Excess non-grain-derived dietaryphosphorus overwhelmed any differences ingrain-derived dietary phytate. However, whendiets were formulated to maintain animal growthbut minimize total dietary phosphorus, such aswhen using a ‘low-ash’ diet that relies more onthe phosphorus provided by the grain compo-nents, benefits in phosphorus and mineral avail-ability, including calcium, zinc and ironavailability, were observed. Trout consumingthese low-ash diets prepared using low-phytategrains excreted between 32% and 44% less phos-phorus than did trout consuming low-ash dietsprepared using ‘normal phytate’ controls. Theseresults indicate potential use of low-phytate grainsin ongoing efforts to reduce the dependence onfishmeal in aquaculture production, and to reduceaquaculture effluent phosphorus.

A follow-up study with trout (Overturf et al.,2003) evaluated mineral availability in experi-mental diets containing barley grain (30% of totaldiet) produced either by the cultivar Harrington(containing ‘normal’ phytate levels of 2.45 mgP/g) or by three of the four Harrington lpa iso-lines listed in Table 8.1. Seed batches used forthis study were produced separately from thoseanalysed in Table 8.1 and were separatelyanalysed for seed phosphorus and other seed con-stituents. Phytate contents were: barley lpa1-1 (M422), 1.15 mg P/g (a phytate reduction of about50%); barley lpa3-1 (M 635), 0.5 mg P/g (a phy-tate reduction of about 80%); barley M 955, <0.1mg P/g (a phytate reduction of >90%). This rep-resented the first animal or human nutritionstudy that evaluated more than one lpa isoline orgenotype of a species in a single study. As seedtotal phosphorus remained fairly constant in the

four lines, the stepwise decrease in seed phytatewas matched by the stepwise increase in seednon-phytate phosphorus, with other seed con-stituents remaining fairly constant.

Sets of isolines like the barley set listed inTable 8.1 provide an experimental model withthe potential to provide a precise measure of theeffects of varying levels of dietary phytate oravailable phosphorus. In fact, Overturf et al.(2003) observed a remarkably linear increase inthe apparent digestibility of dietary calcium infish diets prepared with barley grains thatoccurred parallel to the predicted linear increasein apparent digestibility of dietary phosphorusand linear decreases in grain phytate. In dietsprepared with Harrington (no phytate reduction),M 422 (50% reduction), M 635 (80% reduction)and M 955 (>90% reduction), the apparentdigestibility of calcium was 12.9%, 27.7%, 46.6%and 59.8%, respectively. This confirms the initialfinding of Ertl et al. (1998), and the results ofother animal (Veum et al., 2001) and humannutrition studies (Hambidge et al., 2005), whichindicated a linear, negative relationship betweendietary phytate and calcium nutrition.

Spencer et al. (2000a) demonstrated thatgrowth performance of pigs fed diets consisting oflpa1-1 maize and no supplementary phosphatewas equal to that observed for pigs fed normalphytate maize with supplementary phosphate.Further, when pigs were raised in a commercialfacility, use of lpa1-1 low-phytate grain as a sub-stitute for normal maize resulted in carcasses withless backfat and a higher percentage of ‘lean’meat. This was the first indication that use oflow-phytate types might enhance product quality,in addition to enhancing the management ofphosphorus in animal production. A US patentwas awarded subsequently, describing how theuse of low-phytate maize as a feed substitute fornormal maize reduces cholesterol in eggs(Stilborn et al., 2002). The nutritional mechanismleading to this unpredicted benefit of consump-tion of a low-phytate feed, reduced fat and chol-esterol, is at present unknown.

Human Nutrition Studies EvaluatingLow-Phytate Crops

The greatest interest in seed-derived dietary phy-tate in terms of human nutrition is related to its

122 V. Raboy

impact on mineral nutrition in populations thatrely on grains and legumes as staple foods (Brownand Solomons, 1991). This concern is primarilywith iron and zinc, although dietary phytate canalso have a negative impact on magnesium(Hurrell, 2003; Bohn et al., 2004) and calcium(Hambidge et al., 2005) nutrition. As discussedearlier, dietary phytate may also have positiveroles as an anti-cancer agent, anti-oxidant andinhibitor of renal stone formation (Graf et al.,1987; Shamsuddin et al., 1988; Singh andAgarwal, 2005; Somasundar et al., 2005).However, human nutrition studies conducted todate with lpa crops have solely addressed themineral nutrition question.

The first human nutrition study conductedwith a low-phytate crop evaluated iron absorptionfrom tortillas prepared from wild-type vs. lpa1-1maize (Mendoza et al., 1998). In this clinical-scalestudy, 13 non-anaemic males consumed tortillaslabelled with one of two stable isotopes of iron.Apparent iron absorption was 49% greater (8.2%of intake) with tortillas prepared from lpa1-1maize compared with the wild-type control (5.5%of intake). This result is also interesting as it per-tains to the level of dietary phytate reduction nec-essary to observe mineral nutritional benefits.A study with soy protein (Hurrell et al., 1992)indicated that dietary phytate had to be reducedby 90% or more to observe a benefit in ironabsorption. The reduction in phytate in lpa1-1maize compared with the wild type is about two-thirds, yet an improvement in iron absorptionfrom test meals was observed by Mendoza et al.(1998). The issue of ‘threshold’ levels of reductionin dietary phytate necessary to observe nutri-tional benefits, and the contribution of studiesutilizing lpa crops to this issue, are discussed fur-ther below.

When a similar experiment was conductedwith wild-type and lpa1-1 tortillas fortified with ahigher level of iron (Mendoza et al., 2001), differ-ences in iron absorption reflecting any benefitfrom reduced dietary phytate content were notobserved. This different outcome has beeninterpreted as representing ‘conflicting results’(Drakakaki et al., 2005). However, while a num-ber of factors may have contributed to this lack ofbenefit observed in the Mendoza et al. (2001)follow-up study compared with the originalstudy, such as differences in nutritional status ofthe test subjects, differences in the diet ingredi-ents (higher tannins that are known to have a

negative effect on iron retention) or methods ofpreparation, the most likely factor was that amuch higher level of iron was provided in the testmeals in the later study (4.4 mg/portion vs. 0.93mg/portion). Therefore, rather than representingconflicting results, perhaps the differencebetween the results of Mendoza et al. (1998) andMendoza et al. (2001) simply reflects somethingobserved in the animal-model studies discussedabove. Just as supplementation with phosphoruscan ameliorate low phosphorus availability inanimal feeds prepared with ‘normal phytate’grains and legumes, supplementation or fortifica-tion with iron can ameliorate the negative impactof phytate in human diets. Supplementation rep-resents an effective and established approach toprovision of dietary needs, and can overcomenegative impacts of dietary phytate. However,there are potential advantages to the lpaapproach. This crop improvement approach usesgenetics and breeding to permanently correct theproblem at its source. When lpa crops are used infoods or feeds, a major need for supplementationis removed. Also, consumption of lpa feeds orfoods can result in multiple or ‘global’ benefits inboth phosphorus and mineral nutrition, such asenhanced iron, zinc, calcium and magnesiumnutrition. In contrast, supplementation with asingle mineral such as phosphorus or iron will notcorrect the negative effect of dietary phytate onzinc, calcium or magnesium nutrition (see below).

A rapid ‘in vitro digestion/Caco-2 cell cultureassay’ system was developed for testing the rela-tive bioavailability of iron in food and meals (Yunet al., 2004). For example, dietary ascorbic acidenhances iron absorption, whereas polyphenoliccompounds such as tannins inhibit iron absorp-tion. The Caco-2 model system accurately repro-duced these alternative effects on iron absorptionin humans (Yun et al., 2004). Figure 8.4 illustratesthe results of the in vitro digestion/Caco-2assay when used to evaluate iron availability inwild-type and lpa1-1 maize (R.P. Glahn andV. Raboy, 2006, unpublished data). This assay isspecifically for the amount of ferritin, an iron-binding protein complex that accumulates in theCaco-2 cells, shown to be an accurate measure ofavailable iron. Available iron in wild-type maizewas clearly enhanced by the inclusion of ascorbicacid (compared with cellular ferritin backgroundlevels; Fig. 8.4). However, lpa1-1 maize had avail-able iron levels equivalent to wild-type maize plusascorbic acid, and the addition of ascorbic acid to


lpa1-1 maize further enhanced relative iron avail-ability in a symmetric fashion.

The enhanced relative iron availability oflpa1-1 maize compared with wild-type maize, asassayed by the Caco-2 model (Fig. 8.4), parallelsthe benefit in terms of iron availability of lpa1-1maize observed by Mendoza et al. (1998) in astudy with human subjects, further validating theCaco-2 assay (R.P. Glahn, USDA–ARS, Ithaca,New York, USA, 2006, personal communication).Further, the enhanced iron availability of lpa1-1maize compared with its wild-type control isequivalent to, or greater than, iron bioavailabilityof any of 15 rice genotypes screened with the

Caco-2 assay (Glahn et al., 2002), 20 early-matur-ing tropical maize varieties screened with thisassay (Oikeh et al., 2003) or of grains produced bymaize genetically engineered to co-express soy-bean ferritin and Aspergillus phytase in the maizeendosperm (Drakakaki et al., 2005). Thus the typeof single-gene allelic change in maize lpa1-1appears to have potential for improving the ironbioavailability of this staple food. The set of fourbarley lpa isolines (Table 8.1) were also evaluatedin the same study (Fig. 8.4), although no differ-ence in relative iron bioavailability was observedbetween wild-type and lpa sibling lines. This lackof difference among barley isolines might be dueto high levels of seed tannins compared withmaize, an explanation given for the difference inresults between the two Mendoza et al. (1998,2001) human nutrition studies, or to some otherunknown difference in barley vs. maize grainchemistry.

The first human nutrition study that used lpamaize and addressed the impact of dietary phy-tate on iron (Mendoza et al., 1998) was followedby several similarly designed clinical studies usingsmall (10–20) numbers of volunteer subjects thataddressed the impact on zinc and calcium. In thecase of zinc, two independently developed lpamaize genotypes were compared with theirrespective wild-type isohybrids as controls: maizelpa1-1 grain with a 60% reduction in phytate; and‘NutriDense Low-Phytate’, a maize hybrid pro-ducing grain with an 80% reduction in phytate(Adams et al., 2002; Hambidge et al., 2004).

Previous studies of the impact of dietaryphytate on zinc nutrition indicated that the molarratio of dietary phytate to zinc is critical to zincavailability. This prior work defined a 10:1dietary phytate/zinc molar ratio threshold abovewhich negative impacts of phytate on zinc reten-tion were predicted. The phytate/zinc molarratios measured for the grain produced by thefour genotypes used in these studies were 37:1(wild-type lpa1-1 control), 28:1 (wild-typeNutriDense Low-Phytate control), 17:1 (lpa1-1)and 7:1 (NutriDense Low-Phytate) (Hambidgeet al., 2004). The corresponding values for ‘frac-tional zinc absorption’, measured following con-sumption, by ten volunteers, of test mealsprepared with different stable isotopes of zinc asmarkers for a given genotype, were 0.151 ±0.071 (wild-type lpa1-1 control), 0.135 ± 0.050(wild-type NutriDense Low-Phytate control),

124 V. Raboy

WT WT+

Ascorbicacid

Ipa1-1 Ipa1-1+

Ascorbicacid

10

Cellbaseline

20

Fer

ritin

(ng

) / c

ell p

rote

in (

mg)

30

40

Fig. 8.4. Caco-2 cell culture assay of relative ironavailability in grains produced by wild-type andlow phytic acid (lpa) 1-1 near-isogenic maizehybrids. Relative iron availability was assayed bymeasurement of cellular ferritin formation. Near-isogenic hybrids were produced using wild-type(WT) and lpa1-1 versions of the inbreds A619 andA632. Phytate represented 3.1 mg P/g dry weightin wild-type (WT) grain and 1.3 mg P/g dry weightin low phytic acid (lpa) 1-1 grain. Grains wereprocessed as described (Glahn et al., 2002; Oikehet al., 2003) and analyses indicated thatprocessing did not significantly impact grainphytate levels. Genotypes were tested with andwithout ascorbic acid (200 fM). Values representmeans (n = 5) ± standard errors (P = 0.05).(Unpublished data kindly provided by Raymond P.Glahn, US Department of Agriculture, Ithaca, NewYork, USA.)

0.285 ± 0.042 (lpa1-1) and 0.383 ± 0.066(NutriDense Low-Phytate). Thus, studies of bothzinc and iron availability from lpa genotypes(Mendoza et al., 1998) indicated a linear relation-ship, rather than a critical threshold, betweenphytate/zinc molar ratio and fractional zincabsorption, and between dietary phytate and ironavailability. This is of practical importance tonutrition science and crop improvement whenaddressing issues of mineral nutrition in popula-tions dependent on grains and legumes as staplefoods. For example, it indicates that even moderateheritable reductions in grain or legume seed phy-tate might result in valuable improvements inboth iron and zinc nutrition when these crops areused as basic foods.

Hambidge et al. (2005) evaluated the absorp-tion of calcium from tortilla meals prepared usinggrain produced by the same maize lpa1-1 isoline,compared with its appropriate wild-type control,used in the above zinc nutrition study. Similarclinical-scale experimental methods were used,including stable calcium isotopes as tracers of lpagrain type, and the volunteer subjects were fivehealthy adult women. Mean fractional calciumabsorptions for wild-type or lpa1-1 maize tortillaswere 0.35 ± 0.07 and 0.50 ± 0.03, respectively, astatistically significant difference (P = 0.003).

These clinical-scale and model system studiesprovide important first-generation data on thepotential value, in terms of nutritional enhance-ment, of genetic changes in seed crops, whetheraccomplished through classical genetics or geneticengineering. However, these small-scale (in termsof numbers of participants), short-duration studiesmust be followed by larger-scale (more partici-pants), longer-duration field studies, in which par-ticipants consume diets prepared with wild-typeand lpa seeds in a non-clinical setting. The small-scale, short-duration clinical studies measureacute phenomena, whereas the larger-scalelonger-duration field studies would measureimpacts on health and nutrition of chronic con-sumption of diets differing in phytate levels. Thefirst such study (Mazariegos et al., 2006) found noclear and large difference in zinc absorption, orany statistically significant difference, in healthyGuatemalan children (whose traditional diets relyon maize as a staple food), when consumingeither wild-type or lpa1-1 maize for a 10-weekperiod. However, children consuming the lpa1-1maize compared with children consuming the

wild-type control did display higher fractionalzinc absorption (14%). The difference was notstatistically significant, indicating that the differ-ences in dietary phytate in this context had nogreat impact on zinc absorption. It could alsoreflect other factors, such as the fact that differ-ences in dietary phytate consumed between thetest groups were less than expected due to othersources of phytate in the diet, smaller thanexpected differences in dietary phytate providedby the different maize types and higher thanexpected levels of zinc absorption by test subjects.This highlights the challenge represented by theneed to conduct longer-duration, larger-scalestudies. It is possible that the results of small-scaleclinical studies that indicate a benefit to reduceddietary phytate do not translate into real-worldbenefits or, alternatively, that it is inherently morechallenging to accurately measure differences innutritional status in larger-scale field studies.Whatever the explanation for these differences, itis ultimately essential to prove a benefit underreal-world conditions.

Studies addressing the impact of reductionsin grain or legume phytate in lpa versions of vari-ous crops on magnesium nutrition in humanshave not yet been conducted. However, a recentfinding concerning the distribution of minerals inwhite rice prepared from lpa1-1 rice comparedwith white rice prepared from the wild-typecontrol has relevance to magnesium nutrition. Incereal grains such as rice and wheat, most phy-tate and minerals, including potassium and mag-nesium, are deposited in the germ and outeraleurone layers of the seed, mostly as mixed phy-tate salts. These outer layers are removed duringmilling as the bran fraction. The centralendosperm, which following whole grain millingends up as white rice and white wheat flower,contains very low levels of phosphorus andminerals. It is possible that a mutation that blocksthe ability of seeds to make phytate might alterthe distribution of minerals in the mature seed.In fact, Bryant et al. (2005) found that phospho-rus, potassium and magnesium levels in milledwhite rice prepared from lpa1-1 rice, a mutationthat reduces seed phytate by about 40% (Table8.1), were increased from 25% to 40% comparedwith levels in milled white rice prepared from thewild-type control. Thus, an additional benefit tothe lpa approach may prove to be that lpamutations can favourably alter the distribution of


minerals in grains such as wheat and rice that aremilled prior to consumption.

In contrast to the results of Bryant et al.(2005), an analysis of fractions obtained followingmilling of the wheat lpa mutant Js-12-LPA foundno large differences in phosphorus and mineralcontents between the mutant and the wild-typecontrol (Guttieri et al., 2004). Clearly, more workis needed to evaluate the impact of lpa mutationson grain mineral content and distribution. Both ofthe above studies evaluated only one mutant, andin each case this mutant only conditioned a ‘mod-erate’ block (<50%) in grain phytate. It would beinteresting to conduct a similar analysis of millingfractions obtained from the set of barley lpamutants listed in Table 8.1 that condition blocksin grain phytate accumulation of up to 90%.Milling analysis of a mutant such as barley M955, in which grain phytate is nearly absent,should provide a definitive test of the hypothesisthat blocking grain phytate synthesis can greatlyalter grain mineral distribution. Three additionalstudies (Liu et al., 2004; Ockenden et al., 2004;Joyce et al., 2005) reported the distribution ofphosphorus and minerals in grain fractions fromthe rice, wheat and barley lpa mutants listed inTable 8.1, and found little difference betweenwild-type and lpa fractions. However, these studiestook a different approach to that of Bryant et al.(2005) and Guttieri et al. (2004) in that theyanalysed fractions obtained following dissection ofgrains into two fractions: ‘germ’, which includedthe embryo and scutellum; and ‘rest-of-grain’,which included both the central endosperm andaleurone layer. Therefore, these later studiescould not detect any differences in mineral distri-bution between the aleurone layer and theendosperm, as could the studies analysing millingfractions.

Seed Total Phosphorus andRuminant Nutrition

While dietary phytate is not thought of as amajor issue in ruminant nutrition, there is on-going evaluation of the possible benefits ofreduced dietary phytate, or the use of phytase asa feed supplement, in this production context.For example, one recent study indicated thatphytase use as a feed supplement for ruminants

might improve phosphorus availability (Kincaidet al., 2005). As seed phosphorus chemistry is nota major issue in some ruminant production sys-tems, a new direction might be to develop geneticresources useful for breeding crops with reducedseed total phosphorus. Dairy and beef cattle canoften consume more phosphorus than needed foroptimal production, which, as in non-ruminantproduction, leads to problems in manure man-agement (Volk et al., 2000; Erickson et al., 2002;Rotz et al., 2002; Toor et al., 2005). Heritablereductions in seed total phosphorus from 25% to50% might translate into reductions in dairy andbeef cattle waste phosphorus of similar magni-tude. Breeding efforts using traditional methodsare currently underway to select for ‘reducedseed total phosphorus’ maize lines (Warden andRussell, 2004).

To date, most lpa mutations have greatlyaltered seed phosphorus chemistry, but notgreatly altered seed total phosphorus. In fact, mostlpa mutations so far have been isolated by screeningfor the high seed inorganic phosphate phenotype, aphenotype most pronounced and therefore mostoften identified in a selection, when seeds have nor-mal levels of total phosphorus but greatly reducedlevels of phytate. However, one barley lpa muta-tion, barley lpa1-1, consistently has reduced seedtotal phosphorus compared with wild-type con-trols, typically ranging from 10% to as high as23% (Table 8.1; Dorsch et al., 2003; Ockendenet al., 2004; V. Raboy, 2006, unpublished data). Itis possible that a previously unnoticed class of lpamutations might be those that alter phosphorustransport and impact seed total phosphorus. TheUS Department of Agriculture–AgriculturalResearch Service collection of barley lpa muta-tions is being re-evaluated in this light.

One might also conduct a second-generationgenetic screen for single-gene mutations thatreduce seed total phosphorus but that do notimpact plant phosphorus, such as mutations thatblock the transport of phosphorus from parentplant to progeny seed. One could screen progenyfrom mutagenized populations directly forreduced seed total phosphorus (Fig. 8.5, left,screen no. 1). As is obvious in Fig. 8.5, a ‘lowtotal P’ seed mutant would also be ‘low-phytate’and, as for barley lpa1-1, may represent a valu-able alternative approach to the dietary problemsassociated with phytate. Alternatively, one couldscreen progeny of chemically mutagenized lpa

126 V. Raboy

lines for those mutations that reduce the highinorganic phosphate phenotype of lpa seeds(Fig. 8.5, right, screen no. 2). Mutations or allelesthat reduce the high inorganic phosphate of lpaseeds would represent one type of reversion ofthe phenotype of lpa mutants. Reduced seed inor-ganic phosphate might also reduce the negativeimpact of lpa mutations on yield, as the high inor-ganic phosphate levels in lpa seeds might suppressstarch accumulation, which is so important toyield. This would also represent a type of rever-sion of the lpa phenotype. It is therefore possiblethat in the future genetics resources that reduceseed total phosphorus might be used in combina-tion with lpa alleles to engineer optimal nutri-tional quality for seed crops.

Summary and Future Directions

Substantial progress has been made in the molec-ular biology and genetics of seed phytate. Low-phytate types of soybean and several major graincrops are available for evaluation. Efforts atbreeding high-yielding low-phytate grain with

good agronomic characteristics and stress toler-ance are underway. Future work in the develop-ment of high-yielding low-phytate crops willfocus on continued breeding with currently avail-able genetic resources, but may also involve engi-neering ‘optimized’ lpa crops by directing effortsat selected candidate genes (Stevenson-Pauliket al., 2005) and by using molecular genetics totarget gene expression changes to specific targettissues such as the germ or aleurone layer. In thisway optimal reductions in seed phytate can beachieved while minimizing the impact of changesin gene expression in non-seed tissues and otherwhole-plant processes. For example, when amutation in a myo-inositol phosphate kinaseblocks seed phytate synthesis, it might also nega-tively impact the way roots and shoots respond tostress. By targeting a block in a myo-inositolkinase activity in the seed only, the negativeimpact on vegetative stress response might beavoided.

In the area of crop improvement, futurestudies will also focus on developing geneticresources useful in breeding ‘low seed total P’crops. The most clear-cut application for suchgenotypes is in ruminant production systems suchas dairy and beef production. However, theymight also prove useful in engineering low-phytate crops in which the negative impact onstarch accumulation and yield, perhaps one out-come of the high inorganic phosphate phenotypeof low-phytate types, is greatly reduced.

A growing number of human and animalnutrition studies evaluating lpa crops indicate thatgenetic reductions in crop seed phytate may havebroad benefits in animal and human nutrition.This includes benefits in phosphorus managementin non-ruminant livestock production, improve-ments in product nutritional quality and benefitsin mineral nutrition in human populations thatrely on cereals and legumes as staple foods.Future directions in the human nutrition fieldmight include additional field-scale studies toevaluate the potential value of low-phytate cropsin a non-clinical setting, involving much largernumbers of participants and a longer durationthan typical clinical-scale studies. Each of the ini-tial generation of clinical-scale human nutritionstudies focused on the impact of dietary phytateon a single mineral nutrient. Future studies mighttherefore address mineral nutrition and health inbroader terms, for example by evaluating iron,


Screenno. 1

0

Genotype

Phytate P Inorganic P Other P

Hypotheticallow total P

Hypotheticallow phytic acid /low total P

Low phyticacid

See

d ph

osph

ate

frac

tion

(mg

P/g

)

2

4

Screenno. 2

Normal

Fig. 8.5. Two types of screens for reduced seedtotal phosphorus. Seed phosphorus fractions infour types of grain genotypes: homozygous wild-type or ‘normal’; low phytic acid (lpa); a hypothetical low total P; a hypothetical low phyticacid P/low total P. The dashed arrows indicate thetwo types of genetic screens currently underway:selection for seeds with reduced seed totalphosphorus (screen no. 1) vs. selection forreduced inorganic phosphate in low phytic acidseeds that normally have high inorganicphosphate (screen no. 2).

zinc, calcium and magnesium nutrition simultan-eously in the same subjects.

Future studies might also utilize sets ofisolines that differ greatly in seed phytate to eval-uate the potential positive roles of dietary phytateas an anti-oxidant and anti-cancer agent (Grafet al., 1987; Shamsuddin et al., 1988; Singh andAgarwal, 2005; Somasundar et al., 2005), or as aninhibitor of renal stone formation (Grases et al.,2000). Past studies that addressed the potentialpositive roles of dietary phytate have not movedfar beyond in vitro assays, cell culture studies or

whole-animal models other than the rat (Vucenikand Shamsuddin, 2003). The ability to producesubstantial amounts of grains or legumes thathave large differences in endogenous phytate, butthat are otherwise nearly identical, provides anexcellent model to test the various hypothesesconcerning the potential positive role of dietaryphytate. For example, ready access to sufficientquantities of low-phytate grains and legumeswould permit large-scale, long-duration studiesthat use large animals. Such studies would servewell as a model for human nutrition and health.

128 V. Raboy

References

Adams, C.L., Hambidge, M., Raboy, V., Dorsch, J.A., Sian, L., Westcott, J.L. and Krebs, N.F. (2002) Zinc absorp-tion from a low-phytic acid maize. American Journal of Clinical Nutrition 76, 556–559.

Adeola, O. (2005) Metabolisable energy and amino acid digestibility of high-oil maize, low-phytate maize and low-phytate soybean meal for White Pekin ducks. British Poultry Science 46, 607–614.

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132 V. Raboy

9 Phytase and Inositol Phosphatesin Animal Nutrition: Dietary Manipulationand Phosphorus Excretion by Animals

Xin Gen Lei1 and Jesus M. Porres2

1Department of Animal Science, Morrison Hall 252, Cornell University, Ithaca,NY 14853, USA; 2Departamento de Fisiología, Universidad de Granada,

Granada, Spain


Salts of myo-inositol hexakisphosphate (phytate)represent between 60% and 80% of the totalphosphorus in plant seeds used to feed animals (seeRaboy, Chapter 8, this volume), but simple-stom-ached species such as swine, poultry and fish donot have hydrolytic enzymes in their upper diges-tive tracts to digest phytate in feed. These speciestherefore require dietary supplemental inorganicphosphate to maintain productivity, but the unuti-lized phytate is excreted, resulting in high phos-phorus concentrations in manure that exceed thephosphorus requirements of most crops ( Whalenand Chang, 2001; Adeli et al., 2005). The manurephosphorus therefore accumulates in soil andincreases the risk of phosphorus pollution of waterbodies (see Leytem and Maguire, Chapter 10, thisvolume). The impact of phosphorus pollution fromanimal manures is being exacerbated by the risingglobal demand for meat, which has resulted in ani-mal production being consolidated into largeintensive feeding operations to improve efficiency.In addition to environmental issues, myo-inositolhexakisphosphate chelates divalent metals such ascalcium, zinc and iron (Cheryan, 1980), whichrenders these nutrients unavailable to simple-stomached humans and animals.

To address environmental concerns sur-rounding phosphorus pollution from animal oper-ations, considerable research has been directedtowards manipulating animal diets to improve thedigestibility of phosphorus in grain feed and mini-

mize phosphorus excretion in manures. Thischapter reviews research on manipulation usingphytase, currently the most common strategy toimprove phosphorus efficiency in animal produc-tion. Further details on phytase can be found else-where in this volume (see Greiner, Chapter 6,and Mullaney and Ullah, Chapter 7), and thedevelopment of low-phytate grains, an alterna-tive strategy of dietary manipulation to reducemanure phosphorus, is also discussed (see Raboy,Chapter 8, this volume).

Manipulation of PhosphorusNutrition and Excretion

The commonly used feedstuffs, maize and soy-bean meal, contain ~0.8–1.1% and 1.3–2.2%phytate, respectively. Protein products such assoy isolate, canola, sunflower or cottonseed mealcontain 1.3–5.0% phytate (Han, 1988; Eeckhoutand De Paepe, 1994; Fernández-Quintela et al.,1997; Kasim and Edwards, 1998; Leske andCoon, 1999; Ravindran et al., 1999a; Shen et al.,2005). Bioavailability of phosphorus from thesefeedstuffs, except for those with relatively highintrinsic phytase activity (Eeckhout and DePaepe, 1994; Viveros et al., 2000; Zimmermannet al., 2002; Shen et al., 2005), is <15% for swine(Cromwell, 1992; Weremko et al., 1997) and

15–40% for poultry (Leske and Coon, 1999).Nevertheless, the digestibility of total phosphorusor phytate for a given feed ingredient may beaffected by diet composition, age of the animal,experimental protocol (i.e. faecal vs. ileal digesti-bility), amount of phosphorus present in the dietor endogenous excretion of phosphorus (Shenet al., 2002).

Several strategies have been applied toreduce phosphorus excretion by animals andits accumulation in soil (Fig. 9.1). Meeting theexact nutrient requirements of phosphorus bydifferent species is the first step to reducing theirphosphorus excretion, and digestible phospho-rus levels are more accurate than total phosphoruslevels for that purpose (Poulsen et al., 1999). Phasefeeding with accurate dietary phosphorusallowances is another cost-effective way to reducefaecal phosphorus excretion. High levels of avail-able phosphorus are required during the earlystages of growth and development of the animals,whereas much lower levels are needed at laterstages (Keshavarz, 2000). Therefore, the amountof phosphorus added to the feed can be signifi-cantly reduced across the growing–finishingperiod (Applegate et al., 2003a; Angel et al., 2005).It is also important to recognize the greater effi-ciency of commercial herds or strains in utilizingnutrients (Havenstein et al., 2003) and their abilityto adapt to a moderate deficiency of phosphorus(Yan et al., 2005). Feed supplements such as vita-min D and its analogues or citric acid canimprove phosphorus utilization, thereby reducingits excretion (Boling et al., 2000; Snow et al., 2004).The recent development of low phytate or highavailable phosphorus crops (Spencer et al., 2000;

Veum et al., 2001; see Raboy, Chapter 8, this vol-ume) has provided hope for a simple and sustain-able solution. Commercially available manureamendments like alum can significantly reducesoluble phosphorus in poultry litter (Moore et al.,1999). Best management practices have beendeveloped to prevent diffuse pollution of surfaceand ground waters by agricultural phosphorus(Sharpley et al., 2001). Above all, dietary supple-mentation of a microbial enzyme, phytase, hasproven to be the most effective tool for animalindustry to reduce phosphorus excretion fromanimal waste to comply with the environmentalregulations.

Nutritional Impacts of Phytase

Phytase is a phosphohydrolytic enzyme that initi-ates the stepwise removal of phosphate from myo-inositol hexakisphosphate (see Mullaney andUllah, Chapter 7, this volume). Numerous exper-iments have shown that supplemental micro-bial phytase at 300–1000 units/kg in swine andpoultry diets improves phosphorus bioavailabilityby 10–35% (Lei and Stahl, 2000). Animalresponse to dietary phytase dose monitored bygrowth performance, apparent phosphorusabsorption, plasma inorganic phosphorus con-centration, plasma alkaline phosphatase activity,and bone ash and breaking strength was eitherlinear or curvilinear (Lei et al., 1993a,b;Kornegay and Qian, 1996; Yi et al., 1996;Gentile et al., 2003). The general estimate is thatinclusion of 300–600 phytase units/kg in swineand poultry diets releases 0.8 g of digestible phos-phorus and replaces either 1.0 or 1.3 g of phos-phorus from mono- and dicalcium phosphate,respectively (Ravindran et al., 1995; Yi et al.,1996; Radcliffe and Kornegay, 1998; Esteve-Garcia et al., 2005). Apparently, the inorganicphosphorus equivalence of phytase can beaffected by the physiological status, housing andbehaviour (coprophagy) of the animals (Kemmeet al., 1997a,b), biochemical properties of theenzyme (Augspurger et al., 2003; Applegate et al.,2003b) and the nutritional indices or parameterschosen to estimate phosphorus availability andphytase efficacy (Esteve-Garcia et al., 2005).Because of the acidic pH optimum and greaterpepsin resistance, bacterial phytase (AppA2) is

134 X.G. Lei and J.M. Porres

Reduction of manurephosphorus excretion

Low-phytatecrops

Use ofsupplemental

phytases

Phase feeding

Dietoptimization

Accurate estimate ofphosphorus

requirements

Fig. 9.1. Strategies for reducing manurephosphorus excretion.

three- to fourfold more effective than fungalPhyA phytase in both swine and poultry diets(Augspurger et al., 2003; Applegate et al., 2003b).Supplemental phytase also improves utilizationof other minerals by animals (Lei and Stahl,2000, 2001; Debnath et al., 2005; Lei and Porres,2005) and humans (Sandberg et al., 1996; Porreset al., 2001, 2005). The potential of phytase toimprove iron availability in human foods is ofspecial interest due to the worldwide prevalenceof iron deficiency.

Phosphorus Reduction in Manurefrom Animals Fed Modified Diets

By improving the bioavailability of dietary phy-tate, supplemental phytase significantly reducesphosphorus excretion by animals. The decreasesrange between 10% and 50%, depending ondietary phosphorus concentration, supplementalphytase activity and diet composition ( Jongbloedet al., 1992; Lei et al., 1993a,b; Yi et al., 1996; Liuet al., 1997; Baxter et al., 2003; Applegate et al.,2003b). According to Jongbloed and Lenis(1992), a pig excretes a total of 1.23 kg of phos-phorus during its life cycle. If supplemental phy-tase produces an average of 30% reduction inmanure phosphorus, ~22,000 t of manure phos-phorus annually in the USA from raising market-ing pigs alone (60 million pigs marketed/year inthe USA × 0.37 kg phosphorus/head) would beprevented from entering the environment.

Supplementation of phytase and the use oflow-phytate grains both decrease the amount oftotal phosphorus present in manure-amendedsoils (Maguire et al., 2003). In addition, supple-mental phytase reduces faecal excretion of cal-cium by up to 50% (Fig. 9.2; Lei et al., 1993a)and presumably other minerals as well (Lei et al.,1993c; Stahl et al., 1999). Nevertheless, effects ofphytase and low-phytate grains on water-solublephosphorus excretion remain unclear. Gollanyet al. (2003) reported a 42% reduction in totalmanure phosphorus in pigs fed low-phytatemaize and no differences in manure phosphorussolubility when compared with pigs fed regularmaize. Similar findings were reported byApplegate et al. (2003a) in poultry fed low-phytatemaize or phytase. Wienhold and Miller (2004)did not find significant differences in the distribu-

tion of phosphorus fractions in manure fromswine fed low-phytate or regular maize, whereasPenn et al. (2004) observed considerable reduc-tions in the amount of phosphorus runoff gener-ated under stimulated rainfall from soils amendedwith manure from turkey fed phytase or low-phytate maize compared with those fed the stan-dard diet. Baxter et al. (2003) found lower levelsof total and soluble phosphorus in slurries of pigsfed phytase, low-phytate maize or a combinationof both when compared with those fed a stan-dard diet. However, the dietary treatments ofphytase or low-phytate maize resulted in a higherproportion of dissolved molybdate reactive phos-phorus in the slurries. The environmental effectsof dietary manipulation on phosphorus solubilityand transfer are discussed in detail elsewhere (seeLeytem and Maguire, Chapter 10, this volume).

Functional Site of Phytase in the Animal

Two research groups ( Jongbloed et al., 1992; Yiand Kornegay, 1996) showed the stomach of pigsto be the major site of supplemental Aspergillus nigerPhyA phytase activity. Low activity was detectedin the proximal small intestine, and negligibleamounts in the distal small intestine. Our recentwork with a bacterial phytase Escherichia coliAppA2 (Pagano A.R. et al., 2005, unpublisheddata) has illustrated that stomach is also the mainfunctional site of the enzyme. Because of itsstronger pepsin resistance compared to PhyA(Rodriguez et al., 1999a), pigs fed AppA2 main-tained similar phytase activity in digesta among

Animal Nutrition 135

0

1

2

Faecal phosphorus Faecal calcium

% o

n dr

y ba

sis

Basal+ 750 units/kg

Fig. 9.2. Reduction of faecal phosphorus andcalcium concentrations in pigs fed 750 units ofphytase/kg of maize–soybean meal diet comparedwith those fed only the maize–soybean meal diet.(From Lei et al., 1993a.)

stomach, duodenum and upper jejunum. By thelower jejunum, all supplemental phytase disap-peared in digesta. However, there was a significantinverse relationship between colon phytase activityand dietary phytase supplementation, indicatingthe strong dependence of colonial microbial phy-tases on phytate substrate. Similar findings werereported by Schlemmer et al. (2001). Likewise, thecrop, gizzard and proventriculus are the majorsites of supplemental activity in poultry, and littleactivity is found in the small intestine (Yu et al.,2004). The total inactivation or degradation ofsupplemental phytase in the small intestine ofswine and poultry precludes any adverse effect ofthe enzyme in releasing soluble phosphorus intothe environment upon storage of manure prior tospreading it into the fields. The reduced colonmicrobial phytase activity in animals fed dietarysupplemental phytase will also decrease free or sol-uble phosphorus in the colon and in manure.

Determinants of DietaryPhytase Efficacy

A number of factors can modulate the effective-ness of supplemental phytase in diets for swine orpoultry (Fig. 9.3). The most consistent factor,and probably the strongest, is the dietary cal-

cium/phosphorus ratio. A ratio of 2:1 or wider hasadversely affected phytase function in both swineand poultry, compared with ratios close to 1:1(Fig. 9.4) (Lei et al., 1994; Qian et al., 1997; Liuet al., 1998; Tamim et al., 2004), which is probablydue to the precipitation of calcium phytate.Therefore, a 1.2:1 ratio of calcium/phosphorus isrecommended for phytase-supplemented diets (Liuet al., 1998). Likewise, adding inorganic phospho-rus to phytase-supplemented diets may reduce theefficacy of the enzyme due to product inhibition.

Several groups have demonstrated a positiveeffect of vitamin D derivatives on the utilization oftotal phosphorus, phytate-phosphorus, calcium,zinc and manganese by poultry (e.g. Edwards,1993; Biehl et al., 1995). Vitamin D may act syn-ergistically with phytase on dietary calcium andphosphorus retention in broilers (Biehl et al., 1995;Qian et al., 1997; Snow et al., 2004; Angel et al.,2005), although the effect seems to be more pro-nounced in poultry than in swine (Lei et al., 1994;Biehl and Baker, 1996).

Supplementation of swine or poultry dietswith citric, formic or lactic acid effectivelyimproves daily gain, feed-use efficiency andapparent total tract digestibility of organic mat-ter, ash, phosphorus, calcium and magnesium(Radcliffe et al., 1998; Kemme et al., 1999; Bolinget al., 2000). These organic acids exert their effectby decreasing stomach pH (Radcliffe et al., 1998)


Dietary determinants of phytase efficacy

Ca/P ratioInorganic P

Vitamin Dderivatives

Organicacids

High intrinsicphytase feedingredients

Liquidfeeding

Combinedmicrobialphytases

Low-phytatecrops

Combined feedenzymes

Phytaseproductionsystems

Fig. 9.3. Determinants of dietary phytase efficacy.

and lowering the rate of gastric emptying.Interestingly, addition of these acids in phytase(Fig. 9.5) or vitamin D-supplemented diets pro-duces additive or synergistic effects on the indi-vidual action of these components on nutrientutilization (Han et al., 1998; Kemme et al., 1999;Maguire et al., 2003; Snow et al., 2004).

Augmented Effects of MicrobialPhytase and Other Strategies

Feed ingredients of wheat, barley, oat and theirco-products contain relatively high intrinsic phy-tase activity (Eeckhout and De Paepe, 1994;Viveros et al., 2000; Zimmermann et al., 2002;

Shen et al., 2005). Inclusion of these ingredientsin swine diets with sufficient phytase activity cantherefore supply adequate phosphorus nutritionfor the growing period (Pointillart et al., 1987) orthe entire growing to finishing period (Han et al.,1997). The amount of microbial phytase activityneeded for swine and poultry diets can bereduced by including high intrinsic phytase ingre-dients such as wheat middlings (Han et al., 1998)or barley (Skoglund et al., 1998; Carlson andDamgaard Poulsen, 2003). However, synergisticeffects have not been observed from the combi-nation of various microbial phytases (Stahl et al.,2001, 2004; Augspurger et al., 2003; Gentile et al.,2003). This is intriguing, because phytases withdistinctly different initiation site, substrate affin-ity, pH profile and proteolysis resistance may befunctionally complementary. Appropriate condi-tions for promoting the possible synergistic func-tions of different phytases should be furtherexplored.

Combined supplementation of phytase withother feed enzymes such as carbohydrases or pro-teases has been employed to improve the overallnutrient utilization of animal feeds. The combi-nation of xylanase, glycanase, or glycosidase withphytase showed additive effects on phytatedigestibility (Zyla et al., 1999a), apparent meta-bolizable energy (Ravindran et al., 1999b; Wuet al., 2004) and growth (Zyla et al., 1999b;Juanpere et al., 2005). Fungal acid protease andcellulase promoted phytase-mediated dephospho-rylation of myo-inositol hexakisphosphate in vitro


3

4

5

6

7

Day 0 Day 10 Day 20 Day 30

Pla

sma

phos

phat

e (m

g/dl

)

1:12:1

400

500

Inorganic P (%)Wheat middlings (%)Microbial phytase (units/kg)Citric acid (%)

a

b

a

Wei

ght g

ain

(g/d

ay)

0.2000

015

3000

015

3001.5

Fig. 9.4. Effect of dietary calcium/phosphorusratio on plasma inorganic phosphate concentra-tions of pigs fed 750 units of phytase/kg ofmaize–soybean meal diet. (From Lei et al., 1994.)

Fig. 9.5. Interaction among dietary microbial phytase, plant phytase and citric acid on daily weight gainof pigs. (From Han et al., 1998.) a vs. b: P < 0.05

(Zyla et al., 1995). Feeding swine or poultry withmultiple enzyme preparations (Omogbenigumet al., 2004) improved growth performance anddigestibility of phytate and other components.

Based on growth performance and serumcriteria, Liu et al. (1997) observed that 250 unitsof phytase/kg diet with liquid feeding was aseffective as 500 units/kg with dry feeding of amaize–soybean meal diet. In addition, soakingplus phytase tended to improve phosphorus andcalcium digestibility in comparison with dry feed-ing plus phytase, thus reducing the amount offaecal phosphorus excretion. Skoglund et al.(1998) found that supplementation of microbialphytase to a barley–rapeseed diet and steeping inwhey for 3 h at 40°C reduced faecal phytate con-tent in pigs by 64%. There was a novel applica-tion of liquid feeding in which phytate wasdegraded by phytase from dietary supplement-ation, intrinsic sources of feed ingredients andproduction by the fermenting microbes. Lacticacid was generated from the fermentation of liq-uid feed, and the population of lactic acid bacte-ria in the gastrointestinal environment of the pigwas increased (Canibe and Jensen, 2003; Carlsonand Damgaard Poulsen, 2003).

Low-phytate varieties of maize (Ertl et al.,1998) and barley (Hatzack et al., 2000) have com-parable levels of total phosphorus to the parentallines, but greater bioavailability of phosphorus toswine and poultry (Ertl et al., 1998; Overturf et al.,2003; Thacker et al., 2004). The potential of theselow-phytate varieties for improving feed phos-phorus utilization can be further augmented byphytase supplementation (Yan et al., 2000;Applegate et al., 2003a; Baxter et al., 2003).Meanwhile, transgenic soybean and canola linesthat overexpress the phyA gene of A. niger havebeen developed, and lead to improved utilizationof phosphorus and calcium by poultry (Denbowet al., 1998; Zhang et al., 2000). Onyango et al.(2004) suggested that differences in glycosylationof E. coli phytase among yeast production systemsmight affect the enzyme efficacy. In contrast,Zhang et al. (2000) found no difference in efficacyof A. niger PhyA phytase derived from a microbialor plant (canola) expression system.

Constraints of Phytase Application

Stability and cost are the two major constraintsfor the application of currently available phy-

tases. An ‘ideal’ phytase should be stable for feedpelleting and storage. The high temperaturesused in the pelleting process denature phytaseand thus reduce the enzyme activity in the finalproducts (Wyss et al., 1998; Igbasan et al., 2000).Phytase coating has been developed to counter-act this destructive effect of thermal processing,but does not ensure a complete release of theenzyme from the granules during its transitthrough the gastrointestinal tract of the animal.Spraying of liquid phytase to feed post pelletinghas been used to bypass phytase denaturation bypelleting, but equipment costs are substantial. Itis also challenging to ensure an even distributionof phytase by spraying (Johnston and Southern,2000). The phytase product should be storedunder dark, dry and cool conditions. Refrigera-tion of the enzyme will extend its shelf life, andpowder preparations are usually more stable thanliquid preparations. If possible, phytase should bestored apart from mineral and vitamin premixes.

The economic returns of phytase applicationare determined by the price of the phytase prod-uct and the equipment needed for phytasesupplementation, in relation to the amountsof non-phytate phosphorus, calcium and otherdietary components that can be spared by the useof phytase. Thus, application of phytase is eco-nomically attractive in places where strict regula-tions impose fines to the excess of animal manurephosphorus. Phytase is generally considered safefor handlers or users, and only minor allergicreactions have been described among workers ofa technical centre for large-scale confection ofphytase in powdered form (Baur et al., 2002).

Developing Thermostable Phytases

A thermostable phytase has been isolated fromA. fumigatus. Compared with phytases isolatedfrom A. terreus, Myceliophthora thermophila or A. niger(Pasamontes et al., 1997; Wyss et al., 1998), thisphytase is more thermotolerant. This feature ofthe enzyme can be modulated by specificity ofbuffers used in the heat treatments (Rodriguezet al., 2000a) and appears to be related to its abil-ity to refold after heat denaturation (Wyss et al.,1998). However, there is no major differencein its three-dimensional structure from that ofA. niger PhyA (Xiang et al., 2004).

Developing thermostable enzymes capableof withstanding feed pelleting has been intensively


studied. State of the art for this purpose includesdirected evolution, in which repetitive rounds ofin vitro diversification are conducted and testedusing high throughput screening methodologies.Garrett et al. (2004) optimized the performance ofE. coli AppA phytase by gene site saturation muta-genesis. They generated a library of single sitemutations of the individual amino acid residues ofthe enzyme. After identification of all single pointmutations that enhanced the thermal stability ofthe enzyme, a new screen was designed to dis-criminate between the most stable of the combi-natorial products using a sequential, recursiveaddition protocol. As a result, a final constructtermed Phy9X was developed with significantlyhigher thermostability and gastric resistance thanthe parent AppA enzyme.

Rational design is another powerful tool forthe development of thermostable phytases. Basedon the role of glycosylation on A. niger PhyA sta-bility (Han et al., 1999), Rodriguez et al. (2000b)designed several mutations to add potential glyco-sylation sites in E. coli AppA phytase expressed inthe methylotrophic yeast Pichia pastoris. Two ofthe mutations showed elevated glycosylation.However, the increased glycosylation did notaffect thermostability of the mutated enzymes.Another mutant did not show a higher degree ofglycosylation, but did show a higher thermostabil-ity and improved kinetic properties comparedwith the wild-type enzyme. These changes wereattributed to a higher number of hydrophobicinteractions and disappearance of a disulphidebond between a G helix and GH loop present inthe α-domain of the protein that would improveits flexibility and catalytic properties. The feedingefficacy of this mutant phytase to release phos-phorus from phytate in a maize–soybean diet wastested in a pig trial (Gentile et al., 2003). However,Wyss et al. (1999) did not observe any effect of gly-cosylation on thermostability of phytase.

Based on primary protein sequence analysisof fungal phytases, Lehmann et al. (2000a) chosethe most conserved amino acid residues and con-structed a novel phytase termed consensus phy-tase-1. The synthetic enzyme featured a higheroptimum temperature (16–29°C increase) andunfolding temperature (15–22°C) when com-pared to the parent phytases, but no change intheir catalytic properties. Stabilization of the con-sensus phytase resulted from a combination ofseveral amino acid residues that were locatedmainly in regions with no defined secondary

structure on the surface of the molecule(Lehmann et al., 2000b, 2002). The consensusphytase was effective in releasing phosphorusfrom phytate in diets for swine or poultry(Gentile et al., 2003; Paditz et al., 2004; Esteve-Garcia et al., 2005). In addition, Jermutus et al.(2001) expanded the development of structure-based chimeric enzymes to the replacement of anentire α-helix on the surface of A. terreus phytase.

Enhancing Proteolysis Resistanceof Phytases

Resistance to proteolysis is of importance forphytase to function in the digestive tracts of ani-mals. Given that phytate hydrolysis mainly takesplace in pig stomach or the crop, gizzard andproventriculus of poultry, resistance to pepsin,the main protease in situ, is desirable for efficientphytases. Rodriguez et al. (1999a) found a greaterpepsin resistance of the recombinant AppA phy-tase from E. coli expressed in P. pastoris than thatof the recombinant A. niger PhyA phytase. In fact,the activity of E. coli phytase was elevated afterthe pepsin digestion. Resistance of E. coli AppAto pepsin digestion has been further confirmed byGolovan et al. (2000). Simon and Igbasan (2002)found a higher susceptibility to proteolytic cleav-age of several fungal phytases when compared toa bacterial or consensus phytase. On the otherhand, A. niger PhyA phytase exhibited a lowersusceptibility to proteolysis when compared towheat or yeast phytases (Phillippy, 1999; Matsuiet al., 2000). However, A. fumigatus phytase washighly susceptible to trypsin (Rodriguez et al.,2000a). Kim et al. (2003) described pepsin andtrypsin resistances of a novel phytase isolatedfrom Citrobacter braakii.

Proteolysis resistance of phytase may beenhanced by site-directed mutagenesis that modi-fies cleavage sites of exposed loops susceptible toprotease action (Wyss et al., 1999). Modificationsof E. coli AppA phytase by gene site saturationmutagenesis caused a 3.5-fold enhancement ingastric stability of the enzyme (Garrett et al.,2004). Gastrointestinal carriers may also be usedto help phytase in reducing stomach and smallintestine proteolytic degradation. Haraldsson et al.(2005) reported the use of high phytase-producingstrains of Saccharomyces cerevisiae as the phytase car-riers. The limitation of this approach is the pH


dependence of phytase expression by the yeast,and the narrow pH profile of the phytase activity.This could be partially overcome by the use ofgenetically modified yeast species that producerecombinant phytases active in the acidic milieuof the stomach.

Improving the pH Profile andCatalytic Properties of Phytases

Isolation and cloning of phytases with higher cata-lytic efficiency than the currently commercializedfungal phytases have been a focus of phytaseresearch. AppA phytase isolated from E. coli andexpressed in P. pastoris production system was asefficient as, or superior to, A. niger PhyA phytase(Stahl et al., 2000). A novel E. coli phytase gene(appA2) was isolated from pig colon with a 95%sequence homology to appA, but the expressedphytase in P. pastoris had different biochemicalproperties (Rodriguez et al., 1999b). As men-tioned above, other groups showed the superioreffectiveness of this AppA2 phytase to that of thecommercialized phytases from Peniophora lycii orA. niger in diets for swine or poultry (Augspurgeret al., 2003; Applegate et al., 2003a).

Most of the mutation studies for improvingthe pH profile of phytase have been done usingA. niger or A. fumigatus phytase. As summarized byTomschy et al. (2002), three major approachescan be taken for the purpose:

1. modification of ionizable groups directlyinvolved in substrate specificity or catalysis;2. replacement of amino acid residues in directcontact with residues located in the active orsubstrate specificity site by means of hydrogenbonds or salt bridges;3. alteration of distant charge interactions bymodification of the surface charge of theenzyme.

Tomschy et al. (2002) improved the activity at lowpH of A. fumigatus phytase and a consensus phy-tase. Their approach included decreasing surfacecharge through glycinamidylation and replace-ment of active site residues using site-directedmutagenesis based on sequence alignments andexperimentally determined or homologicallymodelled three-dimensional structures of A. niger

PhyA phytase, P. lycii phytase or A. niger pH 2.5acid phosphatase.

Using molecular modelling of the substratespecificity site of A. niger phytase and sequencecomparisons, Mullaney et al. (2002) successfullydeveloped a recombinant phytase with the substi-tution of lysine by glutamic acid at position 300,which enhanced the specific activity of the enzymeat pH levels between 4 and 5. The authors sug-gested that the residue substitution would lowerthe local electrostatic field attraction for myo-inosi-tol hexakisphosphate at both pH ranges and thusenhance the enzyme kinetics. Recently, our labo-ratory (Kim et al., 2006) produced a shift of opti-mal pH of A. niger PhyA phytase from 5.5 to3.5–4.0 by altering the charges of amino acidresidues in the substrate binding site. The variantshowed a significant enhancement of function inthe stomach of pigs.

By substituting the glutamic acid residuelocated at position 27 by leucine, as in A. terreusphytase, Tomschy et al. (2000a) improved thespecific activity of A. fumigatus phytase withoutchanging its substrate specificity. The improve-ment appeared to be due to a weakening or lossof a hydrogen bond between glutamic acid atposition 27 and the 6-phosphate group of myo-inositol hexakisphosphate, suggesting that prod-uct release would be the rate-limiting step of theA. fumigatus phytase reaction.

In a different set of experiments, Tomschyet al. (2000b) improved the catalytic properties ofA. niger T213 phytase by site-directed mutagenesisof R297Q. A. niger T213 phytase has 12 divergentamino acids from those of A. niger NRRL 3135phytase. Of these divergent amino acids, three arelocated in the vicinity of the active site and weresubstituted by their divergent counterparts ofA. niger NRRL 3135 phytase. The R297Q substitu-tion improved specific activity and pH profile of A.niger T213 phytase to levels comparable to A. nigerNRRL 3135 phytase. As the wild-type A. niger T213phytase had a lower specific activity for myo-inositolhexakisphosphate, but not for other smaller and/orless negatively charged phosphate substrates, theysuggested that product (myo-inositol pentakisphos-phate) release was the rate-limiting step in theenzyme reaction due to interaction of the guani-dino group of arginine in position 297 of A. nigerT213 phytase with one of the phosphate groups ofmyo-inositol pentakisphosphate.


To improve the catalytic properties of thethermostable consensus-1 phytase, Lehmann et al.(2000b) replaced all the divergent amino acidresidues present in the active site of the consensusphytase by those of A. niger NRRL 3135 phytase.The new phytase, termed consensus-7 phytase,featured a major shift in the catalytic propertiesthat were similar to those of A. niger NRRL 3135phytase, thus demonstrating the feasibility ofrational transfer of favourable catalytic proper-ties. However, the active site residues transfercaused a decrease in the unfolding temperatureof consensus-7 phytase compared with consensus-1phytase.

The Search for Efficient PhytaseProduction Systems

Selection of the phytase expression system is cru-cial to ensuring an efficient and affordableenzyme production. The expression system ofchoice may vary with the origin and properties ofthe phytase to be expressed. Because transgenicplants appear to be a cost-effective fermenter forbiological materials, successful attempts to over-express microbial phytases have been madein soybean, wheat, rice, lucerne and canola(Li et al., 1997; Brinch-Pedersen et al., 2000;Ponstein et al., 2002; Ullah et al., 2002; Honget al., 2004). The effectiveness of phytases pro-duced in these plants have been determined (Penet al., 1993; Denbow et al., 1998; Zhang et al.,2000; Hong et al., 2004), but practical problemsinclude public concern over genetically modifiedorganisms and the relatively low thermostabilityof the expressed phytases in plants (Igbasan et al.,2000; Lucca et al., 2001).

Fungal systems from the Aspergillus genera(A. niger, A. oryzae and A. awamori) are employed forphytase production (Mitchell et al., 1997; Martinet al., 2003). Several strategies have been usedto avoid proteolysis associated with Aspergillusexpression systems, including supplementation ofa complex inducing medium with increasing con-centrations of yeast autolysate or additional pro-tein sources like malt extract or casamino acids.Methylotrophic yeast such as P. pastoris orHansenula polymorpha combine several appealingfeatures such as efficient post-translational modifi-

cation, capability of secreting extremely highlevels of recombinant protein, ease of manipula-tion and rapid growth. The enzyme productioncan be greatly enhanced by optimizing fermenta-tion conditions (Mayer et al., 1999; Chen et al.,2004). Rodriguez et al. (1999a,b, 2000a,b)reported efficient heterologous expression ofA. niger, E. coli and A. fumigatus phytase in P. pastorisusing inducible expression driven by the potentalcohol oxidase promoter (AOX1). Lee et al.(2005) expressed the AppA2 phytase using bothinducible and constitutive expression systems,and obtained lower yields in S. cerevisiae andSchizosaccharomyces pombe than in the P. pastorisexpression systems. Never the less, the expressionsystems or hosts had no effect on biochemicalproperties of the recombinant phytases. Thesesimilarities offer flexibility for choosing phytasefermentation systems.

Several bacterial expression systems havebeen studied for phytase expression. To avoidextensive manipulation and purification steps dueto periplasmic expression of recombinant pro-teins by E. coli, Miksch et al. (2002) tested extra-cellular expression of E. coli phytase using asecretion system based on the controlled expres-sion of the kil gene. They analysed major factorssuch as promoter type, host strain and selectionpressure that could affect the level of heterolo-gous expression, and developed an effective fed-batch fermentation strategy. Kerovuo et al. (2000)studied a novel Bacillus expression system anddemonstrated its potential application for effi-cient extracellular phytase production, whereasStahl et al. (2003) attempted extracellular expres-sion of E. coli AppA phytase in Streptomyces lividansand studied changes in its biochemical propertieswhen compared to a glycosylated AppA pro-duced by P. pastoris.

Bacillus subtilis phytase has been expressedin Lactobacillus plantarum 755 (Kerovuo andTynkkynen, 2000). Despite the low expressionand secretion levels, the authors suggested thatculture conditions could be optimized for ahigher yield. In addition, the bacterial straincould be used as an inoculum for fermented plantmaterial or as phytase carrier in combinationwith the probiotic effects inherent to lactic acidbacteria. In general, bacterial expression systemshave a disadvantage compared to plant, fungalor yeast expression systems for their inability to


glycosylate the recombinant protein expressed.Glycosylation has been shown to be importantfor fungal phytase expression (Han and Lei,1999). Phillippy and Mullaney (1997) found thatwhen the A. niger phytase gene was expressedintracellularly in E. coli, the expressed protein wasinactive. Yin et al. (2005) recently described anovel phytase expression system based on silk-worm larvae infected with baculovirus trans-fected with the baculovirus transfer vector pVL

1393 under the control of the polyhedron pro-moter and harbouring the E. coli appA phytasegene. Using this system, the authors obtained aphytase yield of 7710 units/ml hemolymph.

Future Perspectives

Figure 9.6 depicts strategies and paths for devel-oping ‘ideal’ phytases that are catalytic-efficient,


Biotechnology

Catalysisand

pH profile

Thermostability

Development of more efficientproduction systems

Resistance toproteolysis

New phytase geneswith higher intrinsic

activity

Chemical modificationSite-directed mutagenesis

Directed evolutionExchange of active site

Directedevolution

Consensusconcept for

thermostability

Structure-basedchimeric enzymes

Naturallythermostable

enzymes

Rationaldesign

Transgenicplantsand

animals

Fungal systemsAspergillus spp.

Yeast systemsSaccharomyces cerevisiae

Pichia pastorisHansenula polymorpha

Schyzosaccharomyces pombe

Bacterial systemsEscherichia coli

Bacillus spp.Streptomyces lividans

Lactobacillus plantarum

Novel expression systemsBaculovirus-infected

silkworm larvae

High intrinsicresistance toproteolysis

Site-directedmutagenesis

Phytasecarriers

Fig. 9.6. Biotechnology for the development of catalytic-efficient, heat-stable, proteolysis-resistant andeconomical phytases.

protease-resistant, heat-stable and cost-effectivefor different species at various physiologicalstages. The success of this attempt will overcomethe current constraints of phytase application.A wide application of phytase in animal diets,together with other nutritional and environmen-tal measures, will help alleviate or eliminate themanure phosphorus pollution problem world-wide. Transgenic plants overexpressing phytasein leaves and seeds have recently been developedand may offer another economic source of phytasefor animal or human nutrition. Meanwhile, low-phytate grains may provide an appreciable levelof available phosphorus to animals. Transgenicpigs (Golovan et al., 2001) with overexpressedphytase in their saliva require lower levels of

inorganic phosphorus supplements in plant-baseddiets and excrete ~70% less of faecal phosphorusthan the controls. Hopefully, these novel tech-nologies can be combined to effectively improvethe utilization of dietary phytate by animals andminimize environmental phosphorus pollutionfrom their manure.

Acknowledgements

The phytase research in Xin Gen Lei’s labora-tory was funded in part by the CornellBiotechnology Program. Dr Porres works under aresearch contract from Junta de Andalucia,Spain, and Project AGL2002-02905 ALI.


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10 Environmental Implications ofInositol Phosphates in Animal Manures

April B. Leytem1 and Rory O. Maguire2

1United States Department of Agriculture–Agricultural Research Service,Northwest Irrigation and Soils Research Laboratory, 3793 N. 3600 E., Kimberly, ID

83341, USA; 2Crop and Soil Environmental Sciences, Virginia Tech, Box 0404,Blacksburg, VA 24061, USA

Animal production in the USA is valued at morethan $100 billion and has consolidated signifi-cantly during the last 20 years, with a largernumber of animals being produced on anincreasingly smaller land base (Kellogg et al.,2000). Manure generated from animal produc-tion is currently estimated to exceed 335 million tof dry matter per year in the USA, while globalmanure production is estimated at ~13 billion tof dry matter per year (Mullins et al., 2005).Manures contain significant amounts of phospho-rus, with values between 6.7 and 29.1 g P/kg ona dry weight basis reported for several speciesof animals (Barnett, 1994). This phosphorusincludes inorganic and organic forms, with thelatter constituting between 10% and 80% of thetotal (Peperzak et al., 1959; Gerritse and Zugec,1977). Inositol phosphates are one of the primaryorganic phosphorus species found in manures,with myo-inositol hexakisphosphate typicallybeing the most abundant (Peperzak et al., 1959;Barnett, 1994; Turner and Leytem, 2004).

The environmental fate of phosphorus inanimal manures is determined in part by thechemical composition of the phosphorus, yet fewstudies have fully characterized manure phospho-rus and determined the effect of the various phos-phorus compounds on phosphorus behaviour insoil. The various forms of organic phosphorusdiffer in the extent of their sorption when appliedto soils, with myo-inositol hexakisphosphate beingstrongly bound while other organic phosphorus

compounds such as nucleotides, DNA and glu-cose phosphates are more mobile (Celi andBarberis, 2005). Phosphorus applied to soil asmanure may also behave differently from mineralphosphate fertilizer, due to other chemical char-acteristics of the manure. Organic matter inmanure can complex iron and aluminium viaorganic ligands, which decreases the precipitationof inositol phosphates with these metals. It alsocompetes for sorption sites in soil, increasingthe concentration of phosphate in solution(Iyamuremye et al., 1996). Inositol phosphates inmanure can also disperse soil colloids and there-fore increase the potential for particulate phos-phorus transport in runoff (see Celi and Barberis,Chapter 13, this volume). Based on this evidence,more detailed information on the forms of phos-phorus in manures, as well as those manurecharacteristics that influence phosphorus sorp-tion, may shed light on the potential for off-sitelosses of phosphorus from land application ofmanure.

This chapter addresses environmental issuesconcerning phosphorus and inositol phosphatesin animal production. We summarize studies onthe phosphorus composition of manures, includ-ing those using traditional extraction proceduresand the more recent application of nuclear mag-netic resonance (NMR) spectroscopy. Finally, wereview how dietary modification and storagealters the phosphorus composition of manures,and explore the impact of such alterations on

phosphorus solubility in soils and the potentialfor phosphorus transfer in runoff.

Why is Manure Phosphorus anEnvironmental Concern?

Consolidation of animal production can gener-ate regional and farm-scale nutrient surpluseswhere nutrient imports in feed and mineral fer-tilizer exceed nutrient exports in crops and ani-mal products (Sharpley et al., 1994; Sims et al.,1998). These nutrient surpluses can in turnincrease the risk of nutrient loss to the environ-ment and pollution of water bodies (Sharpley,1996; Sims et al., 1998, 2000). Nutrients inmanures can be recycled by application to crop-land, which reduces the need for commercial fer-tilizers. Unfortunately, large amounts of manureproduced in localized areas, coupled with thehigh cost of effective nutrient utilization strate-gies in an unbalanced system, favour manuredisposal via land application in excess of cropnutrient needs, rather than utilizing manure inareas with nutrient deficiencies (Sharpley et al.,1998).

Phosphorus is a particular concern, becauseit can accumulate in soil to concentrations greaterthan those needed for optimum crop production.This is due in part to unfavourable nitrogen/phosphorus ratios in manures relative to theuptake of these nutrients by most crops, whichresults in overapplication of phosphorus whenmanures are applied to meet the nitrogenrequirement of the crop (Mikkelsen, 2000). As aresult, long-term manure application to agricul-tural land leads to soil phosphorus accumulationand greater potential for phosphorus transfer inrunoff to water bodies. This can contribute toeutrophication in freshwater ecosystems, andnumerous examples of water quality impairmentassociated with phosphorus pollution from ani-mal operations now exist (Burkholder andGlasgow, 1997; US Geological Survey, 1999;Boesch et al., 2001). There is therefore an urgentneed to understand and reduce the impact of ani-mal manures on the pollution of water bodies.This demands a mechanistic understanding ofthe behaviour of manure phosphorus in soils andits potential for phosphorus transfer in runoff.Important aspects include the manure character-

istics that determine phosphorus behaviourfollowing land application and the potentialchanges induced by dietary modification.

Phosphorus Composition ofAnimal Manures

Investigation of the dynamics of manure phos-phorus following application to soils requiresinformation on the phosphorus compositionof the manure. One of the earliest studies ofmanure characterization was performed byFunatsu (1908), who used sequential extractiontechniques to fractionate the phosphorus inguano. The procedure involved dilute acid toextract inorganic phosphate, inositol phosphatesand other organic forms, followed by ether andalcohol to extract phospholipids, with the residue(unextracted fraction) being labelled as nucleicacid. Variations of this procedure were subse-quently used by others to characterize manuresfrom pigs fed a variety of feed rations (Rather,1918), poultry and mixed farmyard manure(Ghani, 1941), sheep manure (McAuliffe andPeech, 1949) and fresh manure from horses, cat-tle, sheep, pigs and hens (Kaila, 1948). Organicphosphorus in these studies ranged between 18%and 50% of the total phosphorus, with the acid-soluble organic phosphorus (which typicallyincluded inositol phosphates) constitutingbetween 0% and 86% of the total organic frac-tion.

Peperzak et al. (1959) used a similar sequen-tial extraction procedure to determine the phos-phorus composition of a variety of manures.Total phosphorus concentrations ranged between4 and 30 g P/kg dry weight, with the inorganicfraction constituting 53–95% of total phosphorus(Table 10.1). In this procedure, myo-inositol hexa-kisphosphate was isolated from the acid extractand was found to represent between 1% and22% of total phosphorus, with other acid-solubleorganic phosphorus forms constituting between3% and 44%. The alcohol-soluble fractions weresmall (0.4–1.3%) while residual phosphorus val-ues ranged between 2% and 27% of total phos-phorus. When manures of different ages wereexamined from a stockyard, the general trendwas a decrease in organic phosphorus from 49%to 32% of total phosphorus over 20 years, with a

Environmental Implications in Animal Manures 151

concomitant decrease in myo-inositol hexakispho-sphate from 3.9% to 1.5% of total phosphorus.

Barnett (1994) published the most recentcomprehensive study on organic phosphorus com-pounds in animal manures using conventionalsequential fractionation techniques. Organicphosphorus in a variety of manures was fraction-ated into phospholipids, nucleic acids, acid-solu-ble organic phosphorus, inorganic phosphate andresidual phosphorus. Inorganic phosphate consti-tuted the greatest proportion of the total phospho-rus, followed in descending order of magnitude byresidual phosphorus, acid-soluble organic phos-phorus and small amounts of phospholipids. Inthis study the myo-inositol hexakisphosphate con-tent was not directly measured, but the acid-solu-ble organic phosphorus fraction, which typicallyincludes the inositol phosphates, ranged between7.8% and 53.4% of the total phosphorus.

Interest in the environmental fate of manurephosphorus prompted recent studies to adopt theHedley fractionation (Dou et al., 2000; Sharpleyand Moyer, 2000; Weinhold and Miller, 2004).This procedure was originally developed to assessphosphorus solubility in soil (Hedley et al., 1982)and involves sequential extraction with water,sodium bicarbonate, sodium hydroxide andhydrochloric acid. Phosphorus extracted in waterand bicarbonate is considered readily soluble,while that extracted in sodium hydroxide (assumedto be associated with amorphous iron/aluminiumand organic matter) and hydrochloric acid

(assumed to be calcium phosphates) is consideredpoorly soluble. However, several problems com-promise the suitability of the Hedley fraction-ation for manures. In particular, phosphoruschemistry differs markedly between soils andmanures, being controlled commonly by iron andaluminium oxides and calcium carbonate in soils(Hedley et al., 1982), and by association with cal-cium and magnesium in manures (Cooperbandand Ward Good, 2002).

Turner and Leytem (2004) used solution 31PNMR spectroscopy to unequivocally identifyphosphorus compounds in the various fractions ofthe Hedley extraction scheme as applied to poul-try, swine and cattle manures. Two main groupsof phosphorus compounds were determined withthis procedure: a readily soluble fraction extractedwith water and sodium bicarbonate and a stablefraction extracted with sodium hydroxide andhydrochloric acid. Organic phosphorus in thereadily soluble fraction included DNA, phospho-lipids and simple phosphate monoesters. Organicphosphorus in the stable fraction consisted mainlyof myo-inositol hexakisphosphate. Since there wasconsiderable overlap between the extracts, theauthors recommended a simpler procedure con-sisting of extraction with sodium bicarbonate toremove the readily soluble fraction (which wouldbe most susceptible to transport in runoff),followed by extraction with a solution contain-ing sodium hydroxide and ethylenediaminetetraacetate (EDTA) to recover the more stable

152 A.B. Leytem and R.O. Maguire

Table 10.1. Concentrations of phosphorus compounds in sequential extracts of animal manures. (FromPeperzak et al., 1959.)

myo-Inositol Other acid- Alcohol-

Total PPhosphate hexakisphosphate soluble P soluble P Residual P

Animal (g P/kg) % of total P

Chick 13–23 53–56 NDa 17–44 0.6–1.0 2–27Hen 7–30 54–81 12–22 3–11 0.1–0.6 5–12Sheep 12 63 2 19 0.4 16Sow 11 83 0.6 13 0.5 3Horse 4–7 73–95 1–2 14 0.8 2–20Steer 8–12 60–64 7–10 12–13 1.0 13–19Bull 9 76 0.5 8 0.7–1.0 14Cow 4–7 67–87 1–5 7–25 1.3 3–14Calf 5 62 3 17 0.4–1.3 17

aND = not detected.

fraction. This method gave near-quantitativerecovery of phosphorus from swine and poultrymanure (Turner, 2004; Turner and Leytem,2004).

Solution 31P NMR spectroscopy has beenused to quantify the phosphorus composition of awide variety of manures (Leinweber et al., 1997;Leytem et al., 2004; Maguire et al., 2004; Turner,2004; Turner and Leytem, 2004; McGrath et al.,2005). These studies indicate that manure phos-phorus is predominately inorganic phosphate, fol-lowed in descending order by phosphatemonoesters, phosphate diesters (nucleic acids andphospholipid), pyrophosphates and, in some cases,phosphonates. Concentrations of myo-inositolhexakisphosphate ranged from non-detectable to80% of the total phosphorus in manures from avariety of ruminant (cattle and sheep) and mono-gastric animals (poultry, swine; Table 10.2). Solid-state 31P NMR spectroscopy has also been appliedto manures (e.g. Hunger et al., 2004), but cannotaccurately assess the organic phosphorus fraction.

As demonstrated by both sequential fraction-ation and solution 31P NMR spectroscopy, themyo-inositol hexakisphosphate content of manurescan vary widely, both among and within species(Table 10.2). There are physiological differencesbetween ruminant and monogastric animals thatcan account for these differences. The diets ofmonogastric animals often include large amountsof cereal grains, in which much of the phosphorusoccurs as salts of myo-inositol hexakisphosphate(phytate); for example, approximately two-thirdsof the phosphorus in maize and soybeans is in thisform (see Raboy, Chapter 8, this volume). Asmonogastric animals do not possess ample gutphytase (McCuaig et al., 1972), manures frompoultry and pigs can contain large amounts ofundigested phytate (although see Leytem et al.,2004). In contrast, ruminant animals have thecapacity to hydrolyse inositol phosphates in theirdiet, and manures from animals fed grass orlucerne-based diets contain little phytate.However, there is evidence that for ruminants feda grain-based diet, metal complexation can pre-vent extensive hydrolysis of myo-inositol hexak-isphosphate and allow it to pass through theanimal intact (see Dao, Chapter 11, this volume).

Dietary effects are also evident within agiven species. For example, manure from layinghens fed maize with varying levels of non-phytate

phosphorus, with and without phytase additions,can contain a wide range of myo-inositol hexak-isphosphate concentrations (35–80% of totalphosphorus, whereas manure from broilers fed adiet consisting mainly of barley contains closer to10% of total phosphorus in this form (Table 10.2).This indicates the importance of determiningdietary impacts on the composition of manurephosphorus excreted from the animal to assess thepotential behaviour of manure phosphorus onceapplied on land. Since it has been demonstratedthat inositol phosphates can sorb strongly to soils(see Celi and Barberis, Chapter 13, this volume),changes in the concentration of myo-inositol hexa-kisphosphate in manure could be of concern froman environmental standpoint (discussed later).

Impact of Dietary Manipulationon myo -Inositol Hexakisphosphate

in Manure

As monogastric animals cannot fully utilize phy-tate in cereal grains, mineral phosphate supple-ments are commonly added to their diets toprevent phosphorus deficiency. As describedabove, this increases phosphorus concentrationsin manure and can lead to phosphorus accumu-lation in soils when manure phosphorus isapplied in excess of crop phosphorus removal(Sims et al., 2000).

To address concerns regarding surplus phos-phorus in manure, strategies involving dietarymanipulation are being widely adopted to reducemanure phosphorus concentrations (see Lei andPorres, Chapter 9, this volume). By reducing phos-phorus excretion, manures with nitrogen/ phos-phorus ratios more closely matching the nutrientneeds of crops can be generated, thereby reducingoverapplication of phosphorus and build-up of soilphosphorus. For monogastric animals that have alimited ability to digest phytate, dietary strategiesinclude the isolation of mutant grains that storemost of the total phosphorus in the grain as inor-ganic phosphate and less as phytate (Raboy et al.,2000; Dorsch et al., 2003, see Raboy, Chapter 8,this volume), thereby enhancing phosphorusuptake by the animal and reducing the excretedphosphorus (Spencer et al., 2000; Veum et al., 2002;Jang et al., 2003; Klunzinger et al., 2005). Supple-mentation of animal feeds with microbial phytase is


also used to increase phytate hydrolysis in the gut,thereby enhancing phosphorus utilization by theanimal (Cromwell et al., 1993; Coelho andKornegay, 1996; see Lei and Porres, Chapter 9,this volume). The combination of low-phytate

grains with phytase additions is also utilized tofurther reduce phosphorus excretion.

In addition to reducing the concentrationsof phosphorus in manure, dietary modification isexpected to influence manure phosphorus com-


Table 10.2. Concentrations of phosphorus compounds in extracts of manures from a selection ofanimals determined by solution 31P NMR spectroscopy. (From Leytem et al., 2004, 2005, 2006 andunpublished data; Maguire et al., 2004.)

Total Phosphate Pyro- myo-Inositolphosphorusa Phosphateb monoestersb phosphateb hexakisphosphateb

Manure g P/kg dry wt

Swine manure, 13.46 (97) 13.02 (94) 0.67 (5) 0.13 (1) Trfresh (barley feed)

Swine lagoon 30.00 (99) 29.15 (97) 0.75 (3) 0.09 (<1) NDliquid

Broiler manure 6.36 (99) 4.46 (70) 1.92 (30) ND 0.74 (12)(barley feed)

Broiler manure 15.61 (96) 7.21 (46) 8.19 (53) 0.21 (1) 7.61 (49)(standardmaize diet)

Broiler manure 9.49 (99) 1.22 (28) 8.17 (86) 0.10 (1) 7.62 (80)(maize, low NPPc)

Broiler manure 9.61 (98) 5.33 (56) 4.04 (42) 0.13 (1) 3.39 (35)(maize, low NPP + phytase)

Broiler litter 13.90 (98) 5.71 (41) 8.38 (60) 0.06 (<1) 7.83 (56)(maize, high NPP)

Broiler litter 10.40 (96) 5.05 (49) 5.74 (55) ND 4.88 (47)(maize, high NPP + phytase)

Turkey litter 15.40 (87) 10.90 (71) 6.74 (44) 0.14 (1) 5.09 (33)(maize, high NPP)

Turkey litter 12.80 (94) 8.56 (67) 4.82 (38) 0.14 (1) 3.45 (26)(maize, high NPP + phytase)

Dairy lagoon liquid 8.80 (93) 7.93 (90) 0.82 (9) 0.06 (<1) 0.37 (4.2)Dairy compost 2.50 (98) 2.28 (91) 0.22 (9) 0.004 (<1) 0.03 (1)Beef manure 4.20 (99) 2.51 (60) 1.60 (38) 0.09 (2) 0.34 (8)

(maize-fed)Beef manure 4.10 (83) 2.65 (65) 1.0 (25) 0.25 (6) ND

(pasture-fed)Sheep (barley-fed) 8.45 (91) 5.52 (65) 1.68 (20) 0.41 (5) 0.47 (6)

aValues are total phosphorus extracted by sodium hydroxide and ethylenediaminetetraacetate (EDTA), and values inparentheses are the proportion (%) of the total manure phosphorus determined by microwave digestion.bValues in parentheses are the proportion (%) of the extracted phosphorus.cNPP = non-phytate phosphorus.Tr = trace; ND = not detected.

position, which may have implications for theenvironmental fate of manure phosphorus(Turner et al., 2002). Potentially the greatestimpact of diet modification in monogastric ani-mals on phosphorus forms in manure is likely tobe changes in the amount of phytate excreted,with a corresponding increase in the proportionof the manure phosphorus that occurs as water-extractable phosphate. Thus, as diet modificationreduces the proportion of the manure phospho-rus occurring as myo-inositol hexakisphosphate,the proportion of water-extractable phosphate inthe manure increases as a fraction of total phos-phorus, even though the total phosphorus con-centration may be reduced. This is particularlyevident for poultry manures (Fig. 10.1) and maybe important when manures are applied to landon the basis of phosphorus content, as is nowcommon in several states in the USA.

Feeding low-phytate grains

Mutant grains that contain substantially less phy-tate than the wild-type equivalent that has tradi-tionally been fed to animals (Raboy et al., 2000;Dorsch et al., 2003; see Raboy, Chapter 8, thisvolume) have recently been developed. At pres-ent there are low-phytate varieties of maize, bar-ley and soybean meal that can be used in feedformulations. Low agronomic yields of thesemutant grains have prevented wide adoption, butfuture improvements are likely, and these grains

will be useful for developing strategies to reducephosphorus excretion by monogastric animals.Large reductions in total phosphorus excretioncan be achieved using these grains (Spencer et al.,2000; Li et al., 2001; Veum et al., 2002; Janget al., 2003; see Lei and Porres, Chapter 9, thisvolume), although only a few studies have deter-mined their impact on phosphorus compositionin manure. Toor et al. (2005) reported a decreaseof only 10% in excreted total phosphorus frombroilers fed diets containing normal maize vs.low-phytate maize, although there was a 47%reduction in the amount of myo-inositol hexak-isphosphate excreted by the birds. Baxter et al.(2003) saw the same trend for swine fed low-phy-tate maize; total phosphorus excretion was onlyslightly reduced, but myo-inositol hexakisphos-phate excretion was reduced by almost 50%.

When low-phytate barleys were included inbroiler diets, manure total phosphorus concentra-tions were reduced by 14–24% (Leytem et al.,2006b; Table 10.3). However, myo-inositol hexak-isphosphate concentrations in manures from alldietary treatments constituted only 3–12% of thetotal phosphorus in the manure, even when asmuch as 91% of total phosphorus in the feed wasphytate. This same trend was also reported forswine in a similar study; total phosphorus excretionwas reduced by ~33% when animals were fed low-phytate diets, yet myo-inositol hexakisphosphate wasexcreted only in trace amounts (Leytem et al., 2004;Table 10.3). This indicates that even though mono-gastric animals do not possess sufficient phytase tohydrolyse phytate in the part of the digestive tractwhere phosphorus sorption takes place, the phytateis not necessarily excreted by the animal.

A possible explanation is that barley dietscontain high intrinsic phytase activity (see Leiand Porres, Chapter 9, this volume), which mightlead to phytate hydrolysis in the animal.However, in a study of swine manure from ani-mals fed diets containing wild-type and low-phy-tate maize, which contains little intrinsic phytase,most of the excreted phosphorus (~80% of totalphosphorus) was inorganic phosphate and therewas little difference in the manure fractionsacross dietary treatments (Weinhold and Miller,2004). A more likely explanation, therefore, isthat phytate is hydrolysed in the hindgut by intes-tinal microflora, even though the animals derivelittle nutritional benefit from this process in thelower intestine.


Manure phytate (% total P)

0 20 40 60 80 100Man

ure

wat

er-e

xtra

ctab

le P

(%

tota

l P)

0

20

40

60

80

100

Toor et al. (2005) Maguire et al. (2004) Leytem et al. (2006a)

Fig. 10.1. The effect of phytate concentration onwater-extractable phosphorus in manures frommodified poultry diets. (From Maguire et al., 2004;Toor et al., 2005; Leytem et al., 2006.)

Feeding microbial phytase as asupplement

There are several different types of phytaseenzymes (see Mullaney and Ullah, Chapter 7,this volume), although they all catalyse therelease of phosphate residues from myo-inositolhexakisphosphate. Phytase supplements are nowa common component of animal diets and havebeen successful in reducing phosphorus concen-trations in manures (see Lei and Porres, Chapter9, this volume). However, the effects on manurephosphorus composition and therefore manurephosphorus behaviour in soils are poorly under-stood.

It would be expected that manures fromdiets that included phytase would have less myo-inositol hexakisphosphate than equivalent dietswithout phytase. This was the case in a study ofmanures from swine fed diets with and withoutphytase (Baxter et al., 2003). Concentrations ofmyo-inositol hexakisphosphate in fresh swinemanure were decreased by 2.0–3.9 g P/kg by

adding phytase to the feed. However, duringstorage of manure from the normal diet for 150days, myo-inositol hexakisphosphate as a percent-age of total phosphorus decreased from 15.5% to8.5%, which was attributed to microbial degra-dation. For the phytase-amended diet thedecrease in myo-inositol hexakisphosphate duringstorage was only between 9.1% and 9.8%, indi-cating hydrolysis by the added phytase prior toexcretion (Baxter et al., 2003). Therefore, after150 days of storage, there was no significant dif-ference in myo-inositol hexakisphosphate concen-trations in swine manures from the two diets.

Maguire et al. (2004) grew three flocks ofbroilers and two flocks of turkeys on the samebed of litter using diets that were ‘high’ and ‘low’in non-phytate phosphorus with and withoutphytase additions. Concentrations of myo-inositolhexakisphosphate in both broiler and turkeylitters from diets that included phytase wereconsistently lower than in litters from equiva-lent non-phytase diets (Table 10.4). Inorganicphosphate levels in the broiler and turkey litters


Table 10.3. Phosphorus concentrations in poultry and swine manure fed either a wild-type barley(Copeland and CDC Bold) or mutant barley with reduced amounts of grain phytic acid content (M 422,M 635, M 955). Phosphorus concentrations were determined by extraction in sodium hydroxide andethylenediaminetetraacetate (EDTA) and solution 31P NMR spectroscopy. Means in the same column (foreach animal type) followed by the same letter do not differ significantly (P > 0.05). (From Leytem et al.,2004; Leytem, A.B., Thacher, P.A. and Turner, B.L., 2006, unpublished data.)

NaOH–EDTA extractable P (g P/kg dry wt)

Feed phytate myo-Inositol(% total Phosphate hexakis-

Grain type phosphorus) Total Pa Phosphateb monoestersb,c phosphateb

Poultry (broiler chicks)

Copeland 91 6.36 (99)a 4.46 (70)a 1.92 (30)a 0.74 (12)aM 422 40 4.48 (93)c 3.42 (69)c 1.53 (31)b 0.34 (7)abM 635 37 4.93 (92)bc 3.20 (72)c 1.29 (29)b 0.34 (8)abM 955 <1 5.15 (92)b 3.88 (75)b 1.24 (24)b 0.14 (3)b

Swine (barrows)

CDC Bold 55 13.46 (97)a 13.02 (94)a 0.67 (5)a TrM 422 50 8.55 (95)b 7.77 (86)b 1.08 (12)a TrM 635 26 8.05 (91)b 7.59 (86)b 1.08 (12)a TrM 955 3 8.36 (95)b 7.78 (88)b 0.91 (11)a ND

aValues in parentheses are the proportion (%) of the total manure phosphorus determined by microwave digestion.bValues in parentheses are the proportion (%) of the NaOH–EDTA extracted phosphorus.cValues for phosphate monoesters include myo-inositol hexakisphosphate and other monoesters.Tr, trace; ND, not detected.

Environm

ental Implications in A

nimal M

anures157

Table 10.4. Dietary studies where phytase has been used to reduce the total phosphorus concentrations in poultry manures and the influence on manurephytate content. Means followed by the same letter (within each column and for each study) are not significantly different (P > 0.05).

Manure characteristics (g P/kg dry wt)

Diet, Phytase WSP/total Phytate P/Animal non-phytate P (%) addition Total P WSPa Phytate P ratio total P ratio References

Turkey 0.56 No 17.8a 6.4a 5.09 0.36 0.28 Maguire et al. (2004)Turkey 0.48 Yes 13.5b 6.3a 3.45 0.47 0.26Turkey 0.42 No 11.8c 5.1b 4.89 0.43 0.41Turkey 0.34 Yes 11.0d 5.0b 3.65 0.45 0.33

Broiler 0.36 No 14.1a 4.7a 7.83 0.33 0.56 Maguire et al. (2004)Broiler 0.26 Yes 10.8c 4.2a 4.88 0.39 0.45Broiler 0.29 No 11.7b 2.2b 7.32 0.19 0.63Broiler 0.20 Yes 9.7d 2.6b 5.35 0.27 0.55

Broiler 0.36 No 13.6a 1.1a 7.8 0.08 0.57 McGrath et al. (2005)Broiler 0.26 Yes 10.7bc 1.0a 5.4 0.09 0.50Broiler 0.29 No 11.2b 0.9a 7.3 0.08 0.65Broiler 0.23 Yes 9.6c 0.6a 4.9 0.06 0.50

aWSP = water-soluble phosphate in manure.

were largely unaffected by dietary phytase. Thiswas most likely due to the benefit of decreaseddietary inorganic phosphate supplements beingcancelled out by the increased phytate hydrolysisby dietary phytase.

McGrath et al. (2005) determined myo-inosi-tol hexakisphosphate in litters from broilers fed avariety of diets with and without phytase addi-tion, and found that concentrations were lower inlitter from diets containing phytase than fromdiets without phytase (Table 10.4). Toor et al.(2005) analysed turkey manure and broiler littersamples from diets with and without phytaseusing X-ray absorption near-edge structure spec-troscopy. Although detection of organic phos-phates was difficult using this technique, theauthors concluded that dietary phytase additiondecreased myo-inositol hexakisphosphate concen-trations in manures and litters, and that dical-cium phosphate was the most abundant form ofphosphorus present.

There has been some discussion in the litera-ture as to whether residual dietary phytase willcontinue to hydrolyse myo-inositol hexakisphos-phate in manures following excretion, hencemaking phosphorus more water-soluble. Angelet al. (2005) used combinations of boiling poultryand swine manures, or added antibiotics, to showthat dietary phytase supplementation had noeffect on phytate hydrolysis following excretion.These authors concluded that the ‘increase inwater-soluble phosphorus as a percent of totalphosphorus post excretion is a function of excretamicrobial activity and not dietary phytase addi-tion’ (Angel et al., 2005). McGrath et al. (2005)stored broiler litters generated from diets ‘high’and ‘low’ in phosphorus, with and without phy-tase, at two different moisture contents for 440days. By comparing the interactions of storagetime and moisture, they showed that myo-inositolhexakisphosphate concentrations decreasedthrough time only in litter that was stored ‘wet’.This was unrelated to dietary phytase and wasinstead attributed to enhanced microbial activityin the wet litter (McGrath et al., 2005). Maguireet al. (2006) fed broiler breeders diets ‘high’ and‘low’ in dietary non-phytate phosphorus, withand without phytase. Soluble phosphorus wassimilar in manure from under the feeder as in aclean area, indicating no effect of spilled feedwhether or not it included phytase. However,under the drinker, manure moisture and soluble

phosphorus were higher irrespective of the diet,presumably due to increased microbial activitybreaking down myo-inositol hexakisphosphateinto more soluble forms. The effects of manure-derived phytase in soils are unknown, althoughdiscussion of the interactions of phytase with soilconstituents can be found elsewhere in this vol-ume (see George et al., Chapter 14).

Combining low-phytate grainsand phytase

In addition to research on low-phytate grains orphytase alone, a few studies have investigated acombination of low-phytate grains and phytase.Baxter et al. (2003) reported that such a combin-ation decreased myo-inositol hexakisphosphate infresh swine manures more than either approachindividually (Table 10.5). This trend was alsoseen in broiler litters, in which myo-inositol hexak-isphosphate decreased from 20% of total phos-phorus in a normal maize diet to 12% and 10%in diets containing low-phytate maize and low-phytate maize plus phytase, respectively (Tooret al., 2005; Table 10.5). Other studies combinedphytase and low-phytate grains in poultry dietsand reported reductions of 27–45% of total phos-phorus and 27–49% of water-extractable phos-phate in the litter, although none determinedmyo-inositol hexakisphosphate directly (Applegateet al., 2003; Miles et al., 2003; Penn et al., 2004).

Manure phosphorus compositionand phosphorus solubility in soil

Manipulating the diets of monogastric animalscan have a large impact on the amount of myo-inositol hexakisphosphate excreted from swine,poultry and fish. In addition, storage of manureprior to land application can also influence inosi-tol phosphate concentrations by promotingmicrobial degradation. This raises an importantquestion: Do differences in inositol phosphateconcentrations influence the solubility and poten-tial transport of manure phosphorus to waterbodies following application to soil?

Release of soluble phosphorus from manure-amended soil varies considerably dependingon the source of the manure applied (i.e. animal


species, diets fed, manure handling and storage).This is primarily due to differences in the concen-trations of total and soluble phosphorus in themanure (Sharpley and Moyer, 2000; Kleinmanet al., 2002a,b; Vadas et al., 2004), but may also bedue in part to variability in other physical andchemical properties of the manure. Inorganicphosphate is relatively soluble in soils comparedto myo-inositol hexakisphosphate, which is stronglyretained and unlikely to be lost as soluble phos-phorus in runoff (Anderson et al., 1974; Leytemet al., 2002). Therefore, variability of the phospho-rus composition of manures, either due to differ-ences in species, manure-handling techniques orthrough dietary manipulation, could increasephosphorus transport from land-applied manuresto water bodies (Vadas et al., 2004).

When a variety of manures (swine, dairy andbeef cattle manures that were handled/stored dif-ferently) were incorporated into semiarid calcare-ous soils, there was no significant correlationbetween myo-inositol hexakisphosphate content(ranging between 0% and 8% of total phospho-rus) and soil phosphorus solubility (Leytem andWestermann, 2005; Fig. 10.2a). In this instance,the small amounts of myo-inositol hexakisphos-phate in the manures were probably insufficientto influence phosphorus solubility in the soil.Instead, phosphorus solubility was clearly influ-

enced by the amount of carbon added to the soil(Fig. 10.2b).

When poultry manures were added to asimilar calcareous soil, the amount of myo-inositolhexakisphosphate in the manures, which rangedbetween 35% and 80% of total phosphorus,was strongly and negatively correlated withbicarbonate-extractable soil phosphate, followingmanure application (Fig. 10.3a). Manures wereapplied at the same total phosphorus rate, so thiscorrelation was almost certainly due to thegreater proportion of water-soluble phosphateadded in manure with lower myo-inositol hexak-isphosphate concentrations. However, the rela-tionship was transient, becoming insignificantafter 9 weeks of incubation (Fig. 10.3b). Thisdemonstrates clearly that when manures areapplied on the basis of phosphorus content, theproportion of myo-inositol hexakisphosphate, andtherefore of water-soluble phosphate, has astrong influence on the solubility of the manurephosphorus soon after application.

Extractable phosphate concentrationsincreased between the second and ninth week ofincubation and were correlated with the amountof myo-inositol hexakisphosphate in the manures.In other words, manures with more myo-inositolhexakisphosphate caused greater increases inextractable soil phosphate over time. Analysis of


Table 10.5. Dietary studies utilizing low-phytate grains with and without the addition of phytase and theeffect on manure phytate content. Means followed by the same letter (within column for each study) arenot significantly different at P = 0.05.

WSP/total P Phytate/total PTotal P WSPa ratio ratio

Animal Diet Phytase g P/kg dry weight Reference

Broiler Normal maize No 22.4a 12.6 0.56 20 Toor et al.(2005)

Broiler Low-phytate No 20.1b 13.5 0.67 12maize

Broiler Low-phytate Yes 15.7c 12.0 0.76 10maize

Swine Normal maize No 25.5a 11.9a 0.47 15 Baxter et al.(2003)

Swine Low-phytate No 20.7b 10.8a 0.52 8maize

Swine Low-phytate Yes 15.2c 7.9b 0.52 5maize

aWSP = water-soluble phosphate in manure.

the manure-amended soils immediately followingincorporation (Fig. 10.4a) and after 9 weeks ofincubation (Fig. 10.4b) using solution 31P NMRspectroscopy demonstrated the hydrolysis of myo-inositol hexakisphosphate in the soil, strongly

suggesting that this was responsible for theincrease in extractable phosphate.

Although myo-inositol hexakisphosphate isstrongly bound in soils, microbes in the semiaridcalcareous soil were able to break it down into


Manure carbon/phosphorus ratio

0 20 40 60 80 100 120


0 2 4 6 8

Bic

arbo

nate

-ext

ract

able

P (

mg

P/k

g)

0

10

20

30

40

(a) (b)

r 2 = 0.16; P = 0.43 r 2 = 0.81; P = 0.01

Fig. 10.2. Relationship between bicarbonate-extractable phosphate and (a) manure phytateconcentration and (b) manure carbon/phosphorus ratio for six manures of varying origin added to acalcareous arable soil (Portneuf silt loam) from Idaho, USA, containing 0.75% organic carbon, pH 7.6and 18% clay. (From Leytem and Westermann, 2005.)

2 weeks

r 2 = 0.99; P = 0.002

0 20 40 60 80 1000

4

8

12

9 weeks

r 2 = 0.48; P = 0.193


0 20 40 60 80 100

Bic

arbo

nate

-ext

ract

able

P (

mg

P/k

g)

(a) (b)

Fig. 10.3. Relationship between the phytate concentration in poultry manure and the bicarbonate-extractable phosphate in manure-amended soil following (a) 2 weeks of incubation and (b) 9 weeks ofincubation. The soil was a calcareous arable soil (Portneuf silt loam) from Idaho, USA, containing 0.75%organic carbon, pH 7.6 and 18% clay. (From Leytem et al., 2006.)

inorganic phosphate within a few weeks. It wouldtherefore not be expected to accumulate in thesesoils following successive manure applications.This confirms the evidence for the relativebioavailability of inositol phosphates in calcare-ous soils (Turner et al., 2003) and may explainwhy some contain no detectable phytate (seeTurner, Chapter 12, this volume). In contrast,the same manures applied to an acidic soilshowed no correlation between added manuremyo-inositol hexakisphosphate and extractablesoil phosphate (Mehlich-3 extraction) on any ofthe sampling dates, with only the manure car-bon/phosphorus ratio being correlated to theextractable phosphate concentrations (r2 = 0.84at 2 weeks of incubation; data not shown).

The solubility of phosphorus in manure-amended soils seems to be influenced by thecharacteristics of the manure applied. In the

short term, manures with large concentrations ofmyo-inositol hexakisphosphate can demonstratelower phosphorus solubility on calcareous soils,although this trend does not seem to hold true foracidic soils. However, due to microbial break-down of myo-inositol hexakisphosphate in appliedmanures and concurrent release of soluble phos-phate, these differences are likely to becomeinsignificant over time. Other manure properties,particularly the carbon content, seem to exert alarge influence on phosphorus solubility followingapplication to both calcareous and acidic soils(Leytem et al., 2005), presumably due to stimula-tion of the microbial biomass and fixation ofphosphorus in microbial tissue. This means thatthe addition of manure results in a lower solublephosphorus concentration than would beexpected from mineral phosphate fertilizer appli-cation. It therefore follows that in the long termthe most important factor to consider for landapplication of manures is total phosphorus,rather than the form of the phosphorus applied.

An important impact of manure inositolphosphates on the loss of phosphorus to waterbodies involves erosion and transport of particu-late phosphorus. Erosion can be severe on agri-cultural land and is potentially responsible for themovement of large amounts of inositol phosphatesto water bodies (see McKelvie, Chapter 16, thisvolume). Erosion can be promoted by inositolphosphates in manures due to the dispersion ofsoil colloids following sorption to soil components(see Celi and Barberis, Chapter 13, this volume).There is almost no information on inositol phos-phate transport in particulate material from agri-cultural land, and it is not discussed further here.However, several stereoisomeric forms of inositolhexakisphosphate have been reported from river-ine-suspended solids (Suzumura and Kamatani,1995). More information can be found in adetailed review of organic phosphorus transferfrom soils to water bodies (Turner, 2005).

Dietary Manipulation and theEnvironmental Fate of Manure

Phosphorus

Manures from low-phytate feed

Although the total phosphorus excreted frommonogastric animals fed a variety of low-phytate


(a)

(b)

Chemical shift (ppm)

0246

4.04.55.05.56.06.57.0

4.04.55.05.56.06.57.0

6.3

5.9

5.1 4.7

4.5

Fig. 10.4. Solution 31P nuclear magnetic resonance(NMR) spectra of extracts of a soil amended withpoultry manure (a) immediately following incorpora-tion and (b) after 9 weeks of incubation. The peakat 6.3 ppm is inorganic phosphate, while the otherfour labelled signals are from myo-inositolhexakisphosphate. The spectra demonstrate therelatively rapid hydrolysis of manure-derived myo-inositol hexakisphosphate in soil. (From Leytemet al., 2006.)

grains has been shown to be significantlyreduced, the impacts of these manures on poten-tial phosphorus losses following long-term appli-cation to agricultural land have not been studied.One of the primary reasons for this is the lack ofsufficient quantities of manure needed for field-scale assessments, particularly multi-year projects.Investigation is therefore limited to laboratory-scale studies.

Gollany et al. (2003) showed a 10% reduc-tion in manure phosphorus availability whenmanure from swine fed low-phytate maize-baseddiets vs. normal maize diets was incorporatedinto a silt loam soil. Leytem et al. (2005) incorpo-rated manure from swine fed a variety of low-phytate barley-based diets and found nosignificant relationship between the amount ofmyo-inositol hexakisphosphate added in themanures and bicarbonate-extractable phosphatein soil (Fig. 10.5a). However, as with previousstudies, there was a strong relationship between theamounts of carbon added with the manures andthe bicarbonate-extractable phosphate (Fig. 10.5b).As the amount of phosphorus excreted by ani-mals fed low-phytate grains is reduced, there is acorresponding increase in the manure carbon/phosphorus ratio, which can enhance the stabi-lization of phosphorus in manure-amended soilscompared with soils amended with manures fromnormal grain-based diets. Therefore, even when

applied on the same total phosphorus basis, thereis a potential environmental benefit to feedinglow-phytate grains when the subsequent manuresare land-applied, at least in the short term.

Manures from phytase-amended feed

Studies have consistently shown reductions inmanure total phosphorus and myo-inositol hexak-isphosphate from swine and poultry that havebeen fed diets with phytase, but only when, asrecommended, inorganic phosphate supplemen-tation is reduced to account for enhanced phos-phorus availability due to phytase addition.However, there has been some disagreementover the effect of added phytase on manurewater-extractable phosphate, which is importantbecause it is linked directly to phosphorus lossesin runoff (Maguire et al., 2005a). Dietary phytaseaddition can decrease total manure phosphorusconcentrations by as much as 45% for poultryand 40% for swine (see Lei and Porres, Chapter9, this volume). These reductions are important,as total phosphorus determines build-up ordecline in soil test phosphorus following landapplication of manures. This is particularly truewhere manure is applied on the basis of nitro-gen content – the effects of changes in manure



0 2 4 6 8 10

Bic

arbo

nate

-ext

ract

able

P (

mg

P/k

g)

20

40

60

80

100

Manure carbon/phosphorus ratio

20 40 60 80

r 2 < 0.01; P = 0.51 r 2 = 0.80; P < 0.001

(a) (b)

Fig. 10.5. The relationship between bicarbonate-extractable phosphate and (a) manure phytateconcentration or (b) manure carbon/phosphorus ratio for manures from swine fed low-phytate grain dietsapplied to a calcareous arable soil (Portneuf silt loam) from Idaho, USA, containing 0.75% organiccarbon, pH 7.6 and 18% clay. (From Leytem et al., 2005.)

phosphorus composition are therefore only likelyto become relevant when manure is applied onthe basis of phosphorus content.

Several studies have surface-applied manuresand litters derived from phytase-amended dietsand measured phosphorus in runoff. Smith et al.(2004a) reported that although dietary phytaseadditions decreased the water-extractable phos-phate in swine manure, this had no significanteffect on soluble phosphorus losses in runoff frommanured soils, relative to manure from a non-phytase-amended diet. This was surprisingbecause equivalent weights of manures wereapplied, so manures with smaller concentrationsof water-soluble phosphate (i.e. from phytase-amended diets) were expected to yield less solublephosphate in runoff. In a similar study, however,soluble phosphate concentrations in runoff imme-diately following the application of poultry litterfrom a phytase-amended diet were lower thanfrom soils that received litter from a normal diet(Smith et al., 2004b). Again, manure was appliedon a weight basis and, importantly, the effectbecame insignificant when three consecutive rain-fall events were included. It should be noted thatin both studies the application of alum (alu-minium sulphate) to the litters considerablyreduced soluble phosphate in litter and in runofffollowing litter application to soil. In one study inwhich dietary phytase significantly increasedmanure water-extractable phosphate, Vadas et al.(2004) reported no significant differences in solu-ble phosphate concentrations in runoff betweensoils amended with poultry manures from phytaseand non-phytase-amended diets, even whenmanures were applied at the same total phospho-rus rate.

Using turkey and broiler litters from equiva-lent phytase- and non-phytase-amended diets,Maguire et al. (2004, 2005b) found that dietaryphytase decreased myo-inositol hexakisphosphatein litters, but generally had little effect on manureinorganic phosphate or soluble phosphate lossesin runoff when manures were incorporated intosoil prior to rainfall. This occurred whether litterwas applied on the basis of nitrogen or phospho-rus content. Where more than one runoff eventwas conducted, soluble phosphate lossesdecreased as the number of runoff eventsincreased, and the effects of diet and manurecharacteristics became less significant. These datahighlight the point that the soluble phosphorus in

manure has a greater impact on runoff solublephosphate concentrations in the short term thanin the long term (Penn et al., 2004; Smith et al.,2004b; Maguire et al., 2005b). However, we stillmust consider the fact that long-term land appli-cation of manures results in the accumulation of alarge pool of phosphorus, which may be availablefor release to runoff water over time. The reduc-tion in total manure phosphorus with phytaseadditions has the long-term benefit of reducingtotal phosphorus additions to fields receiving con-tinual nitrogen-based manure applications thatoverapply phosphorus compared to crop needs.

Manures from low-phytate grainsand phytase-amended feeds

As already discussed, combining low-phytategrains and phytase was shown to result in greaterreductions in manure total phosphorus thaneither strategy on its own. It has also been shownto reduce water-extractable phosphate by27–49% (Maguire et al., 2005a). Smith et al.(2004b) reported that adding phytase to poultrydiets containing low-phytate maize led to less sol-uble phosphate in runoff compared to that from anormal diet, but was not different to soluble phos-phate in runoff from diets containing phytase orlow-phytate maize on their own when manureswere surface-applied at the same total phosphorusrate. Penn et al. (2004) observed similar concentra-tions of soluble phosphate in runoff from soilsreceiving surface application of turkey manure(same total phosphorus applied) from normal orlow-phytate maize plus phytase diets. As there areonly a limited number of studies measuring runofffrom soils amended with these manures, it is tooearly to draw firm conclusions. However, the con-sistent reduction in total phosphorus and water-extractable phosphate in the manures suggests aclear benefit in terms of water quality.

Summary

Research to date has shown manure compositionto be heavily dependent on both animal speciesand diet. In particular, differences in feed compo-sition and phytase supplementation mean that


manures from monogastric animals contain awide range of myo-inositol hexakisphosphate con-centrations. However, manure tends to be storedfor long periods of time prior to land application,which allows microbial activity to break down alarge fraction of the myo-inositol hexakisphos-phate. This creates manures that have low myo-inositol hexakisphosphate concentrations whenthey are eventually land-applied. An importantconsequence is that other manure characteristics,such as the carbon/phosphorus ratio, may have agreater influence on subsequent phosphorus solu-bility in the short term than the phosphorus com-position of the manure upon excretion from theanimal. This must be considered when assessingthe effects of dietary manipulation on the envir-onmental impact of manure phosphorus.

When manure is applied to soil, a variety offactors can influence the phosphorus solubilityand the potential for phosphorus transport towater bodies. In the case of surface-appliedmanure, the water-extractable phosphate concen-tration has the greatest influence on soluble phos-phate losses when rainfall immediately followsmanure application. When manures are incorp-orated into soils, other factors control phosphorussolubility and the potential for phosphorus lossesto water bodies. In calcareous soils with loworganic matter contents, phosphorus sorption canbe influenced in the short term by the myo-inositolhexakisphosphate content of the manure, becausemanure with large concentrations of myo-inositolhexakisphosphate lead to small increases in soilphosphate solubility compared with manuresdominated by inorganic phosphate. However, thiseffect is reduced as myo-inositol hexakisphosphateundergoes hydrolysis and contributes to theextractable phosphate pool, at which point otherfactors, such as the manure carbon/phosphorusratio, determine differences in phosphate solubil-ity. In contrast, when manures are applied toacidic soils, there seems to be no influence of myo-inositol hexakisphosphate content on extractablesoil phosphate, and other manure characteristicsmay have a greater influence on phosphorus solu-bility. In situations where phosphorus losses aredominated by soil erosion and particulate phos-phorus losses, the phosphorus concentration inthe soil will overwhelm any influence of theapplied manure phosphorus forms.

Concern has been expressed about thepotential negative environmental implications of

diet alteration on phosphorus losses from manure-amended soils, but given the urgent requirementto reduce total phosphorus concentrations inmanures in areas of high livestock density, dietarymanipulation is overwhelmingly beneficial.Such manipulation may increase the proportionof the manure phosphorus that is soluble in water,but this is likely to have negative environmentalconsequences only when manure is applied on aphosphorus basis and without prolonged storageprior to land application. If manures are appliedon an equivalent weight or nitrogen basis, dietmodification will result in less total phosphorusbeing added to soils and therefore a reduction insoil test phosphorus build-up over time. This inturn decreases the risk of phosphorus transfer towater bodies. In addition, most research indicatesa reduction or no increase in phosphorus losses inrunoff from soils amended with manures frommodified diets compared with normal diets, whenthese are applied on an equivalent phosphorusbasis (surface application or incorporation ofmanures). It therefore seems likely that in mostcases there is no enhanced environmental riskfrom dietary modification and associated changesin manure phosphorus composition.

Future Research Needs

There is an increasing body of research aimed atunderstanding the influence of manure phospho-rus composition on the potential environmentalimpacts related to land application of manure. Atpresent, few studies have determined manurephosphorus composition using techniques such assolution 31P NMR spectroscopy, yet this informa-tion provides valuable insight into the behaviour ofphosphorus in manure after land application andcan help identify the potential risks of modifyingmanures through diet manipulation.

The study of dietary impacts on manurephosphorus composition and subsequent environ-mental risk is becoming more important. Thereare few studies that have detailed the impacts ofaltering animal diets on manure phosphoruscomposition, and these have focused primarily onphosphorus in feeds (i.e. non-phytate phosphoruslevels and the use of phytase). Dietary com-ponents, such as the calcium/phosphorus ratioin feeds, micronutrient additions and carbon


composition of feeds, can alter the manurephosphorus composition and influence water-extractable phosphate, but have not been investi-gated in detail (see also Dao, Chapter 11, thisvolume). There is evidence that calcium andother divalent cations often found in micronutri-ent supplements can bind with myo-inositol hexa-kisphosphate, making both less available duringdigestion (Maenz et al., 1999). Future studiesshould therefore look beyond just dietary phos-phorus in order to understand the extent towhich we can alter the phosphorus compositionin manures and maximize the benefits of dietarymanipulation.

The use of low-phytate grains in animalfeeding operations has received considerableinterest (see Raboy, Chapter 8, this volume) andfurther research will be necessary as new grainsbecome available, especially as these becomeeconomically viable. Low-phytate grains have anadvantage over phytase addition, because they

minimize the interference that dietary inputs(such as calcium and other micronutrients) mayhave on phytate digestion and phytase efficacy.An important drawback at this point to usinglow-phytate grains is the issue of identity preser-vation (ability to keep low-phytate grains separatefrom other grains during processing), which willhopefully be overcome in the future.

Now that modified diets (phytase additions,low-phytate grains and lower phosphorus) arewidely implemented, there is a need for long-termstudies to determine the environmental effects ofmanure application resulting from these diets andthe effects on soil phosphorus forms. There are nolong-term trials studying the effect of land-appliedmanures from low-phytate diets on soil organicmatter, soil phosphorus availability and forms, orphosphorus losses in runoff. Given the importanceof understanding the impact of intensive animaloperations on the phosphorus pollution of waterbodies, such studies are urgently required.


References

Anderson, G., Williams, E.G. and Moir, J.O. (1974) A comparison of the sorption of inorganic orthophosphate andinositol hexaphosphate by six acid soils. Journal of Soil Science 25, 51–62.

Angel, C.R., Powers, W. J., Applegate, T. J., Tamim, N.M. and Christman, M.C. (2005) Influence of phytase onwater-soluble phosphorus in poultry and swine manure. Journal of Environmental Quality 34, 563–571.

Applegate, T. J., Joern, B.C., Nussbaum-Wagler, D.L. and Angel, R. (2003) Water-soluble phosphorus in freshbroiler litter is dependent upon phosphorus concentration fed but not on fungal phytase supplementation.Poultry Science 82, 1024–1029.

Barnett, G.M. (1994) Phosphorus forms in animal manures. Bioresource Technology 49, 139–147.Baxter, C.A., Joern, B.C., Ragland, D., Sands, J.S. and Adeola, O. (2003) Phytase, high-available-phosphorus corn,

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tor dinoflagellate. Journal of Eukaryotic Microbiology 44, 200–205.Celi, L. and Barberis, E. (2005) Abiotic stabilization of organic phosphorus in the environment. In: Turner, B.L.,

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11 Ligand Effects on Inositol PhosphateSolubility and Bioavailability in Animal

Manures

Thanh H. DaoUnited States Department of Agriculture–Agricultural Research Service, BeltsvilleAgricultural Research Center, Room 121, 10300 Baltimore Avenue, Building 306

BARC-EAST, Beltsville, MD 20705, USA


In regions with high concentration of confined ani-mal production operations, continuous manureapplication over many years has resulted in agricul-tural soils that contain high levels of phosphorus(Simard et al., 1995; Frossard et al., 2000; Pautlerand Sims, 2000; Zhang et al., 2002; Lehmann et al.,2005). The build-up increases the potential forphosphorus leaching and transport in runoff tonearby water bodies, with the associated risk ofwater quality impairment (Gerritse and Eksteen,1978; Chardon et al., 1997; Koopmans et al., 2003).A large proportion of the phosphorus in animalmanures can be in the form of inositol phosphates,which occur mainly as salts of myo-inositol hexak-isphosphate (phytate) (Peperzak et al., 1959;Gerritse and Eksteen, 1978; Dao, 2004b; Tooret al., 2005a). As a result, there is considerable inter-est in the identification and behaviour of inositolphosphates in manures and soils.

Other chapters in this volume discuss inosi-tol phosphates in animal nutrition, excretion inmanures and the fate of manure phosphorus inthe environment (see Lei and Porres, Chapter 9,and Leytem and Maguire, Chapter 10, this vol-ume). This chapter examines the processes thatinfluence the solubilization and dephosphoryla-tion of these inositol phosphates in manures. Thisis of particular importance in an environmentalcontext, because these processes influence therelease and bioavailability of inositol phosphatesin soils and water bodies. An in situ ligand-based

enzymatic hydrolysis method has been developedto assess the bioavailability of inositol phosphatesin manure. The procedure illustrates that charac-terization of phosphorus in manures, based on acombination of chemical and biological assays,may more appropriately reflect the availability ofinositol phosphates to organisms and plants. Themild fractionation approach may also revealimportant information about the stability andbiological activity of complexed forms of inositolphosphates, which are often considered to bechemically unreactive in the environment (seeMcKelvie, Chapter 16, this volume).

Assessing the Solubility and Release of Inositol Phosphates in

Animal Manures

Mixed salts of myo-inositol hexakisphosphateaccount for 60–90% of the total phosphorus inseeds of wheat (Triticum aestivum L.), rice (Oryzasativa L.), maize (Zea mays L.), soybean [Glycinemax (L.) Merr.] and mung bean [Vigna radiata (L.)R. Wilczek var. radiata] (Scott and Loewus, 1986;Lott et al., 2000; see Raboy, Chapter 8, this vol-ume). myo-Inositol hexakisphosphate is also foundin leaves of a variety of plants, includingArabidopsis thaliana (L.) Heynh. (Bentsink et al.,2003), Telfairia occidentalis Hook F. (Ladeji et al.,

1995), Euphorbia hirta L. and Launaea taraxacifolia(Wild.) Amin ex C. Jeffrey (Wallace et al., 1998).Diets of domestic livestock and poultry thereforecontain abundant myo-inositol hexakisphosphate.However, swine (Sus scrofa domesticus) and poultry(Gallus gallus domesticus) are monogastric and donot possess sufficient levels of phytase enzymes intheir digestive system to break down inositolhexakisphosphate, which therefore occurs inlarge concentrations in their manures (Zyla et al.,2000; Council for Agricultural Science andTechnology, 2002; Turner, 2004; Toor et al.,2005b; see Leytem and Maguire, Chapter 10,this volume).

In contrast, it is generally thought that inruminant livestock such as cattle (Bos taurus), sheep(Ovis aries) and goats (Capra aegagrus hircus) phytasesproduced by the rumen microflora and phytasesin saliva and intestinal mucosa catalyse thehydrolysis of myo-inositol hexakisphosphate duringthe rumination process, releasing phosphate forassimilation by the animal. However, a numberof recent studies have reported evidence for thepresence of myo-inositol hexakisphosphate inmanure from cattle fed grain-based diets (Dao,2003; Jayasundera et al., 2005; Toor et al., 2005a).Inositol phosphates could escape into the manureof ruminants following reaction with polyvalentcations to form insoluble precipitates. However,the scarcity of information on the speciation ofinositol phosphates in manure and their hydro-lysis to release inorganic phosphate presents amajor challenge to understanding phosphorusdynamics in manure and its fate in the environ-ment following land application.

To assess the composition and solubility ofphosphorus in animal manure, some studieshave adapted fractionation methods developedfor soil phosphorus. Sequential fractionationprocedures were developed to distinguish phos-phorus pools based on the differential solubilityof calcium, aluminium and iron phosphates instrong acids and bases (Chang and Jackson,1957; Hedley et al., 1982; Kuo, 1996). Solid-phase extraction media that included ion-exchange resins or iron oxide-impregnatedpaper were used as anion sinks to measure thelabile phosphorus fraction in soil (van Diestet al., 1960; Abrams and Jarrell, 1992; Chardonet al., 1996; Myers et al., 1999).

However, methods designed for soils maynot be readily applicable to manures. In particu-

lar, manures are composed of partly digestedfeed and are primarily organic, whereas soilshave a mineral phase and an associated complexorganic carbon phase. These matrices requirecaustic chemical treatments to release the differ-ent phosphorus forms associated with soil mineraland organic matter surfaces that exist as a rangeof insoluble inorganic and organic phosphatecompounds in various stages of crystallization(Lindsay, 1979; Graf, 1986).

The possibility that soil phosphorus analyticalmethods do not yield discrete chemical fractionsalso exists. Chemical transformations and transfersbetween phosphorus pools occur as a result of thechoice of solvents, the chemical composition ofthe extractant solution or the harshness of theextraction conditions (Adams and Byrne, 1989;Leinweber et al., 1997; Turner, 2004; Turner andLeytem, 2004; McDowell and Stewart, 2005). Forexample, the concentration of sodium hydroxidein the extracting solution can alter the distributionof phosphate monoesters and diesters in alkalineextracts of animal manures and also influencesspectral resolution in solution 31P nuclear magneticresonance (NMR) spectroscopy (Leinweber et al.,1997; Turner, 2004). The extraction of phosphatesby strong acidic and basic extractants is not lim-ited to single species of phosphate; for example,aluminium phosphate and iron phosphate areboth soluble in acidic extractants. Solubility prod-ucts average 20.3 and 28 for aluminium and ironphosphates, and 32.5 and 96 for aluminium andiron hydroxy-phosphates, respectively (de Haaset al., 2001).

Methods involving the enzymatic dephospho-rylation of phosphorus-containing compoundshave been used to characterize inositol phosphatesand other groups of organic phosphorus inextracts of animal manures (He and Honeycutt,2001; Dao, 2003) and soils (Shand and Smith,1997; Hayes et al., 2000; Turner et al., 2002; Tooret al., 2003; Dao, 2004a; Dao et al., 2005).Phosphatases catalyse chemical reactions thatrelease phosphate from various types of organicphosphorus compounds. That is, when a manuresample or an extract of a manure sample is incu-bated with a specific phosphatase, the release ofinorganic phosphate in the reaction medium indi-cates the presence of a specific type of organicphosphorus compound and its concentration inthe sample. For example, He and Honeycutt(2001) used a modified sequential extraction

170 T.H. Dao

method for soil phosphorus (Hedley et al., 1982)and subjected the various extracts of swine andcattle manure to hydrolysis by acid and alka-line phosphatases, phytases, nuclease P1 andnucleotide pyrophosphatase. They found myo-inos-itol hexakisphosphate-like compounds and smallquantities of phosphate diesters. This methodappears to provide qualitative information on thecomposition and potential bioavailability of themanure organic phosphorus. However, enzymaticmethods have been hampered by the low reactiv-ity of hydrolysable organic phosphorus in extracts,which means that a large proportion of theorganic phosphorus remains uncharacterized. Forexample, in the study described above (He andHoneycutt, 2001), hydrolysable phosphorusaccounted for <50% of the organic phosphorusextracted by water and sodium hydroxide, and<15% of the organic phosphorus extracted bybicarbonate and hydrochloric acid.

In addition, the extent of phosphate releasefrom organic phosphorus substrates may also beaffected by the specificity and purity of the vari-ous enzyme preparations. For example, commer-cial phytase preparations (e.g. phytase fromSigma Chemical Company, St Louis, Missouri,USA) express non-specific acid phosphataseactivity, which can lead to an overestimation ofphosphate release from inositol phosphates com-pared to the use of more highly purified and thusmore specific phytases (Shand and Smith, 1997;Hayes et al., 2000). Ideally, enzyme-based assaysto determine the phytase-hydrolysable phosphateshould employ phytases with high specific activityfor myo-inositol hexakisphosphate.

The fact that relatively small proportions oforganic phosphorus in extracts of manure andsoil can be hydrolysed by phytase has been attrib-uted to the complexation of organic phosphateswith polyvalent cations. Dao (2003) observed thatthe inefficiency of enzymatic methods was notrelated to insufficient levels of enzyme activity orinositol phosphate substrates, but was due in partto the association of inositol phosphates withpolyvalent cations that control their solubility andsusceptibility to dephosphorylation. A similar sit-uation was found in studies of in situ hydrolysis ofphosphate monoesters in soils (Otani and Ae,1999; Dao, 2004a). The phenomenon has impor-tant impacts on the solubility and reactivity ofinositol phosphates in animal manures, and isexplored later.

Inositol Phosphate Reactivitywith Polyvalent Cations

The six phosphate moieties give myo-inositolhexakisphosphate the potential of 12 coordinateligands for complexing cations. It complexesmonovalent cations and stoichiometrically formsmonomeric salts such as sodium phytate.Multivalent cations such as calcium or iron(III)ions can form intramolecular bonds, bridging twoor more phosphate moieties on a single inositolmolecule, resulting in monomeric inositol phos-phates; intermolecular bonding can occur whentwo or more inositol phosphate molecules share acommon multivalent cation and yield polymericcompounds (Vohra et al., 1965; Evans and Pierce,1982; Pierce, 1985; Champagne et al., 1990). Forexample, chemical interactions between myo-inositol hexakisphosphate and cations wereevident in the finding that a high calcium/phosphorus molar ratio in poultry diets interferedwith the effectiveness of dietary phytases. Theformation of insoluble calcium phytate under theintestinal conditions rendered the compoundresistant to enzymatic hydrolysis (Qian et al.,1997; Kornegay, 2001). One would expect simi-lar reactions involving lower-order inositol phos-phates, although the chelation potential isreduced with a decrease in the number of phos-phate moieties on the inositol molecule. Forexample, clinical studies showed that zinc andiron absorption in humans increased with myo-inositol tris-, tetrakis- and pentakisphosphate,compared with the hexakisphosphate form(Sandström and Sandberg, 1992; Sandberg et al.,1999). Therefore, the extent to which polyvalentcations influence the physical state of inositolphosphates depends on three factors: the cation,the cation concentration or molar ratio of cationto inositol phosphate and the interaction withother cations present.

Effects of counterion valencyand solution-phase pH

By varying the molar ratio of cation to myo-inositol hexakisphosphate, it was observedthat dephosphorylation of myo-inositol hexak-isphosphate was not affected by divalent calciumions at a mole fraction of 1:6, in comparison to

Ligand Effects in Animal Manures 171

monovalent sodium ions (Fig. 11.1). Calciumphytate was soluble at low molar ratios of cal-cium to myo-inositol hexakisphosphate-phospho-rus, but dephosphorylation was reduced by 50%when the molar ratio of calcium to myo-inositolhexakisphosphate increased to 6:1 (i.e. a 6:6 cal-cium/phosphate ratio), due to the precipitationof insoluble calcium phytate. Divalent calciumions did not completely inhibit dephosphoryla-tion to the same extent as aluminium and iron,because calcium phytate remains soluble at pH 4(Wise and Gilburt, 1981).

As a strong ligand, myo-inositol hexakisphos-phate had a high affinity for aluminium andiron(III) ions, so dephosphorylation was progres-sively inhibited as the concentrations of thesecations or the mole ratio of cation to myo-inositolhexakisphosphate increased (Fig. 11.1). In alu-minium and iron treatments, phosphate release

decreased by an average of 27% and 32% com-pared with the sodium treatment at 1:6 cation/myo-inositol hexakisphosphate molar ratio or 0.25mM of metal ions at pH 4.5. The phenomenonhas been referred to as an ‘in solution’ sequestra-tion of inositol phosphate; although no visibleprecipitate was observed, the formation of metalchelates sequestered and shielded some of theinositol phosphate from dephosphorylation byphytase (Dao, 2003). More than 80% and 99%inhibition of dephosphorylation was observedwhen molar ratios of aluminium or iron to myo-inositol hexakisphosphate exceeded 3:6 and 6:6,respectively.

myo-Inositol hexakisphosphate is more suscep-tible to dephosphorylation at pH 6 than pH 4.5(Fig. 11.1b). The molecule has 12 ionizable protonsand 6 of them have a pKa ≥ 5.2, while the remain-ing pKa values are <3.2 (Evans and Pierce, 1982).

172 T.H. Dao

1.0

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2:1 Na:IP6-P1:6 Ca:IP6-P3:6 Ca:IP6-P6:6 Ca:IP6-P

2:1 Na:IP6-P1:6 Ca:IP6-P3:6 Ca:IP6-P6:6 Ca:IP6-P

2:1 Na:IP6-P1:6 Fe:IP6-P

3:6 Fe:IP6-P6:6 Fe:IP6-P

2:1 Na:IP6-P1:6 AI:IP6-P3:6 AI:IP6-P6:6 AI:IP6-P2:1 Na:IP6-P

1:6 AI:IP6-P3:6 AI:IP6-P6:6 AI:IP6-P

2:1 Na:IP6-P1:6 Fe:IP6-P3:6 Fe:IP6-P6:6 Fe:IP6-P

Time (min) Time (min)

PO

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e co

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Fig. 11.1. Short-term kinetics of the dephosphorylation of myo-inositol hexakisphosphate as affectedby polyvalent counterion concentrations and solution-phase pH (a=pH 4.5; b=pH 6.0). (Reprinted fromDao, 2003.)

As solution pH increases, more metal hydroxidespecies, such as M(OH)2+, M(OH)2+ and M(OH)3(where M = aluminium or iron) and consequentlyless Al(H2O)6

3+ or Fe(H2O)63+ aquo-metal ions,

would exist in solution. The formation of moreamorphous Al(OH)3 and Fe(OH)3 would reducethe effective concentrations of aluminium and ironthat can react with myo-inositol hexakisphosphate.Also, more inositol phosphate remains in dissoci-ated forms and is susceptible to dephosphorylationat pH 6.0, compared to pH 4.5. Therefore, theinhibition of dephosphorylation by aluminium andiron has been attributed primarily to chelation,resulting in sterically hindered forms of myo-inositolhexakisphosphate.

The presence and persistence of myo-inositol hexakisphosphate

in ruminant excreta

Similar reactions to those described above affectedmyo-inositol hexakisphosphate added to liquiddairy manure suspensions (Dao, 2003). Solution-phase phosphate increased with the addition ofAspergillus ficuum phytase, but this was reduced insamples with added polyvalent cations. The addedphytases hydrolysed an average of 22 mmol P/kgdairy manure solids (native phytase activity in themanure was deactivated by steam sterilization).This demonstrated the presence of soluble, fullyionized and uncomplexed myo-inositol hexakispho-sphate in dairy manure that was amenable tohydrolysis by added phytase. It did not representall the inositol phosphates in dairy manure,although the phytase-hydrolysable phosphorus(PHP) assay was subsequently improved with theuse of a combination of phytases and polydentateligands (Dao, 2004a,b) (see below).

On the basis of these results it was postulatedthat these sequestration reactions are mechanismsby which inositol phosphates in feed grain persistduring passage through the animal, leading toexcretion in the faeces of dairy cattle (Dao, 2003).In the animal digestive tract and in manure, myo-inositol hexakisphosphate and lower-order inositolphosphates likely interact with polyvalent cations,primarily calcium, aluminium, iron and magne-sium, and to a lesser extent with manganese, zincand copper, as these micronutrients are added tothe feeds as mineral supplements for nutritional

and health considerations (National ResearchCouncil, 2001).

The combination of the rapid rate of feedpassage through high-productivity dairy cattleand the reduced susceptibility of complexed inos-itol phosphate to dephosphorylation by complex-ation with dietary polyvalent cations results inthe persistence and excretion of complexed myo-inositol hexakisphosphate in dairy manure. Thus,while rumen microflora or intestinal mucosa phy-tases are theoretically capable of hydrolysingmyo-inositol hexakisphosphate in feed, this doesnot appear to happen in practical feeding regi-mens. Manure from dairy and beef cattle fedgrain-based diets can therefore contain consider-able amounts of myo-inositol hexakisphosphate( Jayasundera et al., 2005; Toor et al., 2005a) and,as for monogastric animals, presents an environ-mental concern.

Characterizing the Relative Stabilityof Inorganic Phosphate and Inositol

Phosphates in Animal Manure

Metal chelate stability

The reactivity of inositol phosphates and theinhibitory effects of polyvalent cations on thedephosphorylation of inositol phosphates meanthat chelation can significantly affect the physical,chemical and biological processes controlling thesolubilization and bioavailability of inositol phos-phates. Chelating agents or ligands are complexorganic anions that have multiple functionalgroups sharing pairs of electrons with a centrallylocated cation. The cations that are commonlyassociated with inositol phosphates are transi-tional metal ions, such as iron(III), zinc and cop-per(II). It is not within the scope of this chapter toextensively discuss coordination chemistry, butthe salient concept essential to the understandingof the extractability and exchangeability of insol-uble inositol phosphates in manure and soils isthe relative strength of metal chelates.

The formation and stability of chelatingagents and cation complexes have been expressedas an equilibrium constant or the ratio of activi-ties of the cation–ligand complex and the dissoci-ated cation and ligand (Anderegg, 1971; Martelland Smith, 1974; Lindsay, 1979). The general


case of ionic equilibria between a cation and achelating ligand can be expressed as

n nM L MLD+ (11.1)

where n = 1, 2, 3, . . ., i. A formation constant forthe MLn species is defined as

[ ] [ ][ ]

K M LML

nn

ML n=

In systems of multidentate ligands such as myo-inositol hexakisphosphate and polycarboxylate ligands (e.g. 1,2-cyclohexanediamine tetraacetatediaminocyclohexane tetraacetate (CDTA) and eth-ylenediaminetetraacetate (EDTA)), the formation ofhydrogen complexes also accompanies the cationchelation process in which one or more of thedonor atoms of the ligand are linked to a proton.The multidentate ligand forms protonated species:

H L HL H K H Lp2D D Dg+

As such, the protonation equilibria must also bedefined as

pH L H LpD+ (11.2)

yielding stepwise protonation constants:

[ ] [ ][ ]

K H H LH L

1p

p

p=

-

Combining Equation (11.1) and Equation (11.2),the overall stability constant of the cation chelateis expressed as

[ ] [ ] [ ][ ]

KM H LMH L

p np n

ML n= (11.3)

A comparison of the likelihood of a cation–ligandcomplex to remain intact can be made betweenpairs of cations and ligands and to determinewhether an exchange between two specific lig-ands would occur. An example of calculations ofcharge concentrations and exchange betweenmyo-inositol hexakisphosphate and polycarboxy-late ligands can be found elsewhere (Dao, 2004b).Phase diagrams of equilibrium relationshipsbetween cations and ligands as a function of pHand oxidation–reduction potential can provideuseful guidelines for developing extracting condi-tions that favour the solubilization and exchangeof phosphate and organic inositol phosphateanions with comparable or stronger ligands. Theconsiderable body of literature on the role oforganic acids in the exchange with sorbed phos-phate has been reviewed (Mu et al., 1995; Joneset al., 2003).

Ligand exchange

Organic anions and plant root exudates have longbeen implicated in the acquisition of phosphorusby plants grown in phosphorus-deficient soils (Aeet al., 1990; Hinsinger, 2001; Ryan et al., 2001;Jones et al., 2003). Excreted organic anions com-prise a variety of products of the citric acid cycle,bearing one or more –COO− functional groupsthat exchange with phosphate anions and chelatethe counterions. For example, piscidic acid (p-hydroxybenzyl tartaric acid) and its p-methoxy-benzyl derivative are released in the rhizosphereof pigeon pea [Cajanus cajan (L.) Millsp.], allowingthe legume to utilize iron-bound phosphate.

In soil, commonly identified low-molecularweight organic anions include formate, acetate,propionate, oxalate and citrate, which are formedduring microbial metabolism and decomposition ofplant residues (Rovira and McDougall, 1967). Soilorganic matter contributes aliphatic acids, phenols,phenolic acids, fulvic and humic substances(Ritchie et al., 1982; Freche et al., 1992; Haynes andMokolobate, 2001). These humic matter ligandsare large and have multiple functional groups,binding cations and releasing soluble phosphorusto the soil solution. Animal manure and wastewateralso contain large amounts of organic matter thatlikely generate complex ligands (Ma et al., 2001).Mechanisms of phosphate and inositol phosphatesolubilization in liquid manures may include ligandexchange, complexation of counterions and solubi-lization (Earl et al., 1979; Jones and Darrah, 1994;Kirk et al., 1999).

Organic ligands have been shown to inhibitthe precipitation of di-, tri- and octacalcium phos-phates or hydroxyapatite, forming phospho–citratecomplexes that inhibit the precipitation reaction indomestic wastewaters (Sharma et al., 1992; House,1999; van der Houwen and Valsami-Jones, 2001).The exchangeability of ligands thus forms the basisfor the development of an approach to enhancingthe efficiency of enzymatic methods to determinebioavailability and solubility of phosphate andinositol phosphates in manures (Dao, 2004b).

Effects of ligand/phosphorus ratioson phytase activity

Owing to the ability of one ligand to participatein exchange reactions with another ligand, adetailed study of the effects of polydentate ligands

174 T.H. Dao

on the dephosphorylation of myo-inositol hexak-isphosphate was conducted in the presence of cal-cium, aluminium and iron at levels that inhibitedthe enzymatic dephosphorylation reaction.Polydentate ligands of different sizes and chargecharacteristics included CDTA, diethylene tri-aminepentaacetate (DTPA), EDTA, oxalic acid(ethanedioic acid) and phthalic acid (1,2-ben-zenedicarboxylic acid). They were assessed fortheir ability to decouple complexed myo-inositolhexakisphosphate, thus allowing enzyme bindingand dephosphorylation to occur.

Effects of ligand/myo-inositolhexakisphosphate molar ratio

It was first established that EDTA and the otherligands did not appear to interfere with thehydrolytic activity of phytases at molar concentra-tions up to 6.1 and 22.6 mM for EDTA andphthalate, respectively, when no polyvalent cationwas present (Figs 11.2a and 11.3a). However,there were contradictory reports of organic lig-ands inhibiting the activity of phytases and broad-spectrum phosphatases. Polyvalent anions such asarsenate, molybdate, L-tartrate and phosphatehave been found to inhibit the activity of acidphosphatases, while EDTA, oxalate and citratesuppressed yeast (Saccharomyces cerevisiae) phytasesbut did not affect the activity of wheat (T. aestivumL.) bran phytases (Nayini and Markasis, 1984,1986). However, the presence of proteins, ligandsand inositol phosphates can interfere with colori-metric measurements of phosphate and underesti-mate the rate of reaction and extent ofdephosphorylation of inositol phosphates (Shandand Smith, 1997; He et al., 1998; Dao, 2003).Therefore, the apparent contradictory evidencemay have been an artefact of the analyticalmethodology at ligand concentrations above criti-cal levels.

Adding ligands to a mixture of myo-inositolhexakisphosphate and aluminium reversed theinhibitory effect of aluminium ions on thedephosphorylation of the inositol phosphate.When aluminium concentrations exceeded 0.75mM, the extent of this phenomenon was inthe following order: phthalate = oxalate < DPTA< EDTA = CDTA. The ligands CDTA andEDTA were able to completely reverse the inhi-bition of inositol phosphate hydrolysis at all alu-minium concentrations up to 1.5 mM. At pH

4.5, CDTA and EDTA were present primarily asdivalent H2CDTA2− and trivalent HEDTA3−,respectively. They were both tetradentate andwere apparently comparable in reactivity andability to decouple counterion and myo-inositolhexakisphosphate for phytases to hydrolyse thelatter compound.

The ligands were not as effective in revers-ing the inhibitory effects of polyvalent cations atpH 6. Overall, EDTA was much more effectivethan CDTA, particularly at aluminium concen-trations >0.75 mM (Fig. 11.3). In fact, EDTAwas the only ligand that reversed the inhibitionof the dephosphorylation reaction when the aluminium concentration was 1.5 mM. Theseresults indicated that charge concentration was amajor factor in countering the inhibition ofdephosphorylation by the cation (see later).Charge concentrations for EDTA4− and thechelating ability exceeded those of CDTA,which theoretically existed as H2CDTA2− andHCDTA3−. Furthermore, aluminium existed asboth Al(OH)2+ and Al(OH)3 (aq) at the higherpH. Amorphous metal hydroxides sorbed andshielded myo-inositol hexakisphosphate fromdephosphorylation. Meanwhile, oxalate, com-monly used to chelate iron and aluminium, wasonly able to partially reverse the inhibition ofdephosphorylation at the 0.25 mM aluminiumtreatment and oxalate/myo-inositol hexakisphos-phate molar ratios ≥3:1 (Fig. 11.3). Oxalate wascompletely ineffective at higher aluminium con-centrations and was even less effective at pH 6,where it should exist as the fully dissociated con-jugate base.

Effects of ligand/myo-inositolhexakisphosphate charge concentration ratio

The polycarboxylate ligands under study pos-sessed two (oxalate and phthalate), four (CDTAand EDTA) or five (DTPA) carboxylate func-tional groups. Increasing charge concentrationsreversed the inhibitory effect of polyvalentcations (Figs 11.2 and 11.3). The susceptibility tohydrolysis increased linearly with ligand/myo-inositol hexakisphosphate charge concentrationratios between 1 and 4. The tetra- and pentaden-tate EDTA, CDTA and DTPA were most ableto dissociate aluminium and myo-inositol hexak-isphosphate, and increased the dephosphoryla-tion of the dissociated anion.


Excess charge was needed to decouple andmobilize the complexed myo-inositol hexakisphos-phate, and to attain the ligand/myo-inositol hexak-isphosphate charge concentration ratios between 1and 4, equivalent molar concentration ratios hadto be between 1.5- and 12-fold that of myo-inositol

hexakisphosphate, except in the case of phthalate.However, there must be an upper limit to increas-ing charge concentration needed to overcome theinhibitory effect of polyvalent counterions, becausemolar concentrations should eventually reach levels that would precipitate the enzyme.

176 T.H. Dao

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1.0

0.8

0.4

0.6

0.2

0.0

1 2 3 4 5 6 1 2 3 4 5 6

Ligand/IP6 mole ratio at pH 6Ligand/IP6 mole ratio at pH 4.5

Sol

utio

n-ph

ase

PO

4−P

rel

ativ

e co

ncen

trat

ion

(C/C

0)

IP6 alone+ CDTA+ DTPA+ EDTA+ Na oxalate+ Na phthalate

(a) (b)

B

A

C

D

Fig. 11.2. Effect of polycarboxylate ligands, ligand/myo-inositol hexakisphosphate mole ratios, andsolution pH on the dephosphorylation of inositol hexakisphosphate by Aspergillus ficuum phytase atfour levels of aluminium counterion a=pH 4.5; b=pH 6.0. (A = 0; B = 0.25; C = 0.75; and D = 1.5 mM).(Reprinted from Dao, 2004b.)

Efficacy of polycarboxylate ligands in dairymanure suspensions

An exchange between ligands and myo-inositolhexakisphosphate anions also took place when bothcompounds were added together to dairy manuresuspensions, which enhanced the enzymaticdephosphorylation of the added organic phos-

phate. Increases in the concentration of aluminiumand iron in the manure suspension predictablyreduced dephosphorylation of myo-inositol hexak-isphosphate. The ligands EDTA, CDTA and, tosome extent, DTPA reversed the inhibitory effectof aluminium and iron(III) ions at cation/myo-inositol hexakisphosphate and ligand/myo-inositol


1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1.0

0.8

0.6

0.4

0.2

0.0

1 2 3 4 0 5 10 15 20 25

Ligand/IP6 charge ratio Equivalent ligand/IP6 mole ratio

IP6 alone+ CDTA+ DTPA+ EDTA+ Na oxalate+ Na phthalate

Sol

utio

n-ph

ase

PO

4−P

rel

ativ

e co

ncen

trat

ion

(C/C

0−1)

A

B

C

D

Fig. 11.3. Effect of organic ligands, ligand/myo-inositol hexakisphosphate charge ratios and equivalentmolar ratios on the dephosphorylation of myo-inositol hexakisphosphate by Aspergillus ficuum phytaseat four levels of aluminium counterion (A = 0; B = 0.25; C = 0.75; and D = 1.5 mM), at a solution pH of4.5. (Reprinted from Dao, 2004b.)

hexakisphosphate molar ratios ≤3:1 and aniron/myo-inositol hexakisphosphate molar ratio of6:1 [1.5 mM iron(III)]. Meanwhile, phthalate andoxalate had even less or no effect on removing theinhibitory effect of aluminium or iron in manuresuspensions compared to simple buffered solutions.

The dephosphorylation of added or nativeinositol phosphates was indistinguishable. Inmanure suspensions amended with only exoge-nous myo-inositol hexakisphosphate, in theabsence of added polyvalent counterions, CDTAand EDTA addition increased the release ofphosphate over and above the added myo-inositolhexakisphosphate. This suggested that these lig-ands increased the solubilization and dephospho-rylation of complexed organic phosphorus nativeto the manure and not merely the dissolution ofinorganic phosphates; that is, there was nodetectable effect of the ligands alone in una-mended manure beyond that released as EDTA-exchangeable phosphorus. The results imply thatthe addition of polycarboxylate organic ligands tothe assay mixture allows a more complete meas-urement of soluble and complexed inositol phos-phate in animal manures.

Differentiating pools of phosphate andinositol phosphates in animal manure

Phosphorus exists in a number of chemical formsthat depend upon feed composition, extent ofmineral supplementation in the feed, feed intakeand absorption efficiency as well as the externalphysical conditions upon excretion. A mild lig-and-based fractionation assay has been developedto differentiate bioavailable phosphate and phos-phate monoesters, including inositol phosphatesin manure (Dao, 2003; Dao et al., 2006) and soils(Dao, 2004a; Dao et al., 2005), into pools thatreflect their potential solubilization. The task iscritical to the accurate assessment of the fate ofmanure phosphorus in the environment, includ-ing the contribution of manure-derived inositolphosphates to eutrophication.

Fractions of manure phosphorus that aremeasured in the PHP assay include: (i) water-extractable phosphate; (ii) an EDTA-exchange-able phosphate pool that was not previouslyextracted by water alone; (iii) a water-extractableorganic phosphorus pool that is hydrolysable byphytase; (iv) an EDTA-exchangeable organicphosphorus pool that is hydrolysable by phytase;

and (v) a residual pool that is not extractable bywater or ligands.

Water-extractable phosphorus fraction. In animalmanures, water-extractable phosphorus includesmainly soluble phosphate and small quantities ofdissolved organic phosphorus species such asphosphate monoesters and nucleotides (Gerritseand Eksteen, 1978; Dou et al., 2000; He andHoneycutt, 2001; Turner and Leytem, 2004).Enzymatic assays for organic phosphorus-containing compounds rely on the detection ofreleased phosphate; hence the determination ofwater-extractable phosphate is essential to theaccuracy of the methods and the effectiveness ofenzymatic methods as a quantitative measure-ment tool for organic phosphorus. Althoughwater-extractable phosphorus would appear to bea simple measurement, much variability and ana-lytical artefacts exist in current methods, resultingin contradictory observations. For example,extraction periods (1–24 h or longer) and solu-tion/solid ratios (10:1 to 200:1) vary widely.Changes in solution chemistry and abiotic factors, such as temperature, pH, oxidation–reduction potential or changes in solution/solidratio, can alter the water-extractable phosphorusconcentration based on the impact on the solubil-ity of mineral phosphate species in manures.Inadequate knowledge of the wastewater chem-istry, coordination chemistry and their effects onphosphate dissolution in complex manure mix-tures may explain in part the observed variabilityin analytical protocols and in water-extractablephosphorus results.

Another common error in the determinationof water-extractable phosphorus is to overesti-mate the dissolved phosphate pool by not recog-nizing the ever-present biological processes.Concurrent phosphate-generating processes suchas hydrolysis of organic phosphorus forms bymanure phytases or phosphatases increases water-extractable phosphate concentrations duringextraction. Therefore, water-extractable phos-phate concentrations determined using longerequilibration periods would also include somephosphate released by enzymatic hydrolysis oforganic phosphorus (Dao et al., 2006). A compar-ison of time series measurements of the liquidphase of undiluted samples and batch-dilutedsamples were made to illustrate the pitfalls ofthese laboratory estimates. These observationsare reminiscent of the misinterpretation of sorp-

178 T.H. Dao

tion and transformation processes affecting labilereactive organic chemicals in microbially activesystems (Dao et al., 1982; Scott et al., 1982; Daoand Lavy, 1987). Common conditions that cancontribute to high apparent water-extractablephosphorus include heat-drying, freeze-drying ofliquid manure or extended equilibration periodsthat promote concurrent production of a com-mon end product by multiple phosphate-generat-ing pathways, including artificially enhancedbiological degradation during measurement.

Using a simple set of standardized protocolsto minimize the latter artefact (a solid/water ratioof 1:100, w/v; 1 h equilibration), water-extractablephosphorus averaged about 15.9% ± 14.8% ofmanure total phosphorus, with a median value of9.9% in a case study of 107 manure samples col-lected from dairy farms located across five states ofthe north-east USA (Lugo-Ospina et al., 2005).Water-extractable phosphorus in freshly collectedmanure also averaged about 37 mmol P/kg or12.9% of the total phosphorus (Dao and Daniel,2002) and both sets of results suggested that mostof the phosphorus in dairy manure was associatedwith the particulate phase.

LIGAND-EXCHANGEABLE INORGANIC PHOSPHATE.Tetra- or pentadentate ligands can be used toinduce an exchange between the added ligand andinorganic and organic phosphates and to deter-mine sorbed and precipitated inorganic phosphatesthat were not previously extracted by water alone.An additional fraction, ranging between 6.3% and22.9% of the total phosphorus was extracted from107 samples of dairy manure (Dao et al., 2006).The ligand forms coordination complexes with the

released cations, preventing re-precipitation ofmineral phosphates and maintaining the releasedphosphorus in the solution phase. The immediatemeasurements of phosphate concentrations in theEDTA extract yielded mainly a measure of com-plexed inorganic phosphates in the manures,because no exogenous phytases were used andhydrolytic enzyme kinetics were found to be slowunder the experimental conditions. The sum of cal-cium and magnesium ions in the EDTA-extractingsolution was correlated to the EDTA-exchangeablephosphorus concentration (Table 11.1). This wasnot unexpected, because high levels of mineral cal-cium are added to the diet of lactating dairy cattle(i.e. 0.37%).

PHYTASE-HYDROLYSABLE PHOSPHORUS. Addingphytases to manure suspensions resulted in furtherincreases in phosphate that were not previouslyextracted by water alone. The net PHP content ofall samples of the manure set averaged 32.2% ±15.6% of manure total phosphorus (as determinedusing a non-specific phytase). These results sug-gest the presence of organic phosphorus substratesthat include myo-inositol hexakisphosphate andother inositol phosphate monoesters in dairymanure collected from farms across five north-eastern states of the USA. Although spilled feedsand bedding materials mixed with the manurescould contribute myo-inositol hexakisphosphate tothe mixtures, previous work has shown that faecesand reconstituted dairy manure, prepared fromfreshly collected faeces and urine, also had a dis-tinct PHP fraction (Dao, 2004b).

EDTA-EXCHANGEABLE PHYTASE-HYDROLYSABLE

PHOSPHORUS. Using both ligands and enzymes in


Table 11.1. Binary relationships between extractable calcium, magnesium and water-extractable andbioavailable phosphorus fractions in 107 manure suspensions collected from dairy farms across fivestates of the north-east USA. (Adapted from Dao et al., 2006.)

Y Variable X Variable (mol/kg)

EDTA-extractable Phosphorus fraction (mol/kg) EDTA-extractable calcium calcium and magnesium

Water-extractable phosphorus y = 1.70 x + 0.033 y = 1.27x + 0.020r 2 = 0.396 r 2 = 0.460

EDTAa-extractable phosphorus y = 0.684x − 0.222x2 − 0.007 y = 0.488x − 0.112x2 − 0.035r 2 = 0.788 r 2 = 0.799

Phytase-hydrolysable phosphorus y = 0.194x − 0.194x2 + 0.037 y = 0.088x − 0.011x2 + 0.047r 2 = 0.432 r 2 = 0.380

aEDTA = ethylenediaminetetraacetate.

the extracting solution yielded the greatest con-centration of phosphorus among any of the previ-ous assays, because the inhibitory effect ofpolyvalent cations that shielded complexed myo-inositol hexakisphosphate from hydrolysis wasremoved by the ligands.

It must be recognized that the discussion isfocused on EDTA, but there is no question thatother polydentate ligands, e.g. CDTA, can beequally or more efficient in the ligand-exchangeprocess. However, the theoretical basis forimprovement in the efficiency of enzymaticmethods for characterizing phosphorus pools inanimal manure and soils remains the same. Inaddition, using analytical methods appropriatefor the chemical species in each fraction is criticalto the proper classification and mass balance ofphosphorus forms in the manure sample (Dao,2003; Dao et al., 2006).

It is important to note that commercial phy-tase preparations used in the studies describedearlier are also active against a number of phos-phate monoester and diester substrates, in addi-tion to myo-inositol hexakisphosphate (McKelvieet al., 1995; Shand and Smith, 1997; Hayes et al.,2000; Turner et al., 2002). Use of a more specificphytase preparation might confirm the identityof the derivatives of feed inositol phosphatesin animal excreta. However, this would notimprove the knowledge of the potential biologi-cal availability of organic phosphorus present inmanure. From the perspective of agricultural andenvironmental sustainability, the critical issue isthat animal manure contains a great deal ofbioavailable organic phosphorus (i.e. compoundsthat can be hydrolysed by phosphatases). Thismeans that the knowledge of all potentiallybioavailable phosphorus forms better reflects themagnitude of the threat of manure phosphorusto water quality following transport in runoff toaquatic environments.

Temporal changes in biologicallyavailable phosphorus

The ligand-based PHP assay defined earlier pro-vides a relative scale of chemical and biologicalstability of phosphorus in manures. Phosphoruspools with greater stability, whether the sub-strates are inorganic phosphates, inositol phos-phates or other organic phosphates, may becomebiologically available over longer time scales in

soil. This was observed in a case study of thepotential for dissolution and solubilization ofimmobilized phosphorus in soils treated withadditives to reduce soluble phosphorus in soils(Dao et al., 2005). A freshly prepared ironhydroxide additive reduced water-extractablephosphorus in soil by 90% over a period of 16weeks. In addition, a plant-available phosphatefraction (Mehlich-3 extractant) was also reducedin iron-treated soils, and both fractions remainedunchanged up to 16 weeks.

The ligand-based PHP assay, on the otherhand, revealed a completely different picture ofthe internal changes in phosphorus pools andshowed that the effect of the iron additive wastransitory (Table 11.2). The inorganic EDTA-extractable phosphorus fraction and the PHP poolwere being remobilized, reaching initial soil levelsto nullify the phosphorus-immobilizing action ofthe iron additives by about the fourth week follow-ing soil treatment. It also appeared that the mobi-lization of the PHP pool occurred as a sequentialmultistage process. The processes of solubilizationand desorption of bioavailable phosphorus in una-mended and Fe(OH)3-amended soils were bestdescribed by single or sequential double exponen-tial kinetic equations. This behaviour would beconsistent with the fact that these substances canrange from meta-stable non-ordered to semi-crystalline physical states (Lindsay, 1979; Zhanget al., 1992; Arai and Sparks, 2002).

After 16 weeks, the total pool of potentiallybioavailable phosphorus had returned to, orexceeded, the initial concentration in untreatedsoils, reaching an average 28% ± 1.0% and 56% ±1.7% of total phosphorus content in the Thurmontand Burch soils, respectively. In comparison,Mehlich-3 phosphorus concentrations represented16% ± 0.4% and 20% ± 0.3% of soil total phos-phorus. Thus, the PHP assay can detect changes inthe susceptibility of soil phosphorus inorganic andorganic pools to transformations that traditionalsoil test methods do not account for.

Conclusions

In areas of intensive animal production, agricul-tural soils have become enriched with phospho-rus because of years of continuous manureapplications. Organic phosphorus is an important

180 T.H. Dao

aspect of this problem, because it represents aconsiderable proportion of the phosphorus inplant residues and animal manures and presentsa threat to aquatic systems following its transferin runoff from soils to water bodies. Speciation ofinositol phosphates in environmental samples ispoorly understood, which hampers the under-standing of the environmental behaviour andtransformations of inositol phosphates. It is there-fore difficult to develop comprehensive strategiesfor managing excess nutrients in animal agricul-tural production systems.

This chapter reviewed factors affecting thesolubilization and dephosphorylation of dis-solved and insoluble complexes of myo-inositolhexakisphosphate in animal manures. An in situligand-based enzymatic method was described,which provides insight into the biological stabil-ity of inositol phosphates in manures. Selectedpolydentate ligands and fungal phytases candifferentiate various phosphorus pools thatcontribute to the solution-phase phosphate con-

centration in animal manures and manure-amended soils.

The biological environment, including theactivity of extracellular phytases, is hard to con-trol and varies widely across soils, agroecosys-tems and climatic regions. Yet it plays a keyrole in regulating the fate of inositol phosphatesin the environment. The inclusion of biologicaland biochemical mechanisms in the methodol-ogy for assessing manure-derived inositol phos-phates may therefore reflect more accuratelytheir availability to microorganisms and plants.Moreover, these mechanisms can reveal theunderlying potential for their dephosphoryla-tion and release of phosphate in the long term.Further studies on the role of manure chemistryand ligands in controlling inositol phosphatesolubilization and mobility will improve ourunderstanding of the linkage between water-soluble, exchangeable and hydrolysableforms, and their mobility in soils and the widerenvironment.


Table 11.2. Kinetic models describing temporal changes in water-extractable and bioavailablephosphorus fractions in unamended soil and soils amended with iron hydroxide. (Adapted from Daoet al., 2005.)

Soils

Phosphorus fraction Amendment Thurmont gravelly loam Burch sandy loam

Water-extractable Unamended y = a + b(exp(−cx)), z y = a + b(exp(−cx)),phosphorus where a = 9.1, b = 12.4, where a = 9.8, b = 15.3,

c = 0.989 c = 0.434+ Fe(OH)3 Not fitted Not fitted

EDTAa-exchangeable Unamended y = a + b(1 − exp(−cx)), y = a + b(1 + (c exp(−dx) −phosphorus where a = 109.6, d exp(−cx))/(d − c)),

b = 12.5, c = 0.0792 where a = 63.9, b = 34.1,c = 0.1931, d = 0.1931

+ Fe(OH)3 y = a + b(1 − exp(−cx)), y = a + b(1 − exp(−cx)),where a = −38.9, where a = 328.6, b = 117.5, c = 0.391 b = 187.1, c = 0.068

Phytase-hydrolysable Unamended y = a + b(1 + (c exp(−dx) − y = a + b(1 + (c exp(−dx) −phosphorus d exp(−cx))/(d − c)), d exp(−cx))/(d − c)),

where a = −4293.8, where a = 172.9, b = 4658.6, c = 2.785, b = 49.7, c = 0.3862, d = 2.7823 d = 0.3862

+ Fe(OH)3 y = a + b(1 − exp(−cx)), y = a(1 − exp(−bx)),where a = −378.4, where a = 165.4, b = 620.5, c = 0.604 b = 0.250

aEDTA = ethylenediaminetetraacetate.

References

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12 Inositol Phosphates in Soil:Amounts, Forms and Significance of thePhosphorylated Inositol Stereoisomers

Benjamin L. TurnerSmithsonian Tropical Research Institute, Apartado 0843-03092, Balboa, Ancón,

Republic of Panama


Inositol phosphates are abundant in soil, yet theirorigin, dynamics and ecological function remainlargely unknown. Given the importance of phos-phorus in agriculture and the environment, it isremarkable that so little is known about one of themost prevalent forms of soil organic phosphorus.In particular, several inositol phosphates that arecommon in soil occur nowhere else in nature.

Inositol phosphates were first reported in soilmore than 60 years ago (Dyer et al., 1940; Yoshida,1940) and their quantitative importance soonbecame apparent. Numerous reports in the follow-ing decades, including the pioneering work ofDennis Cosgrove and George Anderson, docu-mented the amounts and forms of inositol phos-phates in soil. However, there has been littleadditional research since Michael L’Annunziata’sstudies of inositol phosphate stereochemistry in themid-1970s (see L’Annunziata, Chapter 4, this vol-ume). This has been due in part to the widespreadadoption of solution 31P nuclear magnetic reso-nance (NMR) spectroscopy as the method ofchoice for the analysis of soil organic phosphorus,which conventionally provides data on phosphatemonoesters as a broad functional group ratherthan as individual compounds such as the inositolphosphates. However, recent advances in method-ology mean that detailed information on soil inos-itol phosphates can now be obtained usingstandard NMR procedures (Turner et al., 2003b;Turner and Richardson, 2004), which should rein-vigorate research on the inositol phosphates in soil.

Other chapters in this volume deal specifi-cally with the reactions, mobility, hydrolysis andbioavailability of inositol phosphates in the envi-ronment. This chapter addresses specifically theamounts, forms and functions of inositol phos-phates in soil. Emphasis is placed on the phos-phorylated stereoisomers, some of which havenever been detected in biological tissue. Thechapter builds on a recent review of inositol phos-phates in the environment (Turner et al., 2002)and the reader is also referred to a series of otherreviews of soil organic phosphorus that includeinformation on the inositol phosphates (Anderson,1967; Cosgrove, 1967; Halstead and McKercher,1975; Dalal, 1977; Anderson, 1980; Cosgrove,1980; Harrison, 1987; Stewart and Tiessen, 1987;Magid et al., 1996; Condron et al., 2005).

Amounts of Inositol Phosphatesin Soil

There is a considerable body of data on inositolphosphates in soil, some of which is summarizedin Table 12.1. From this it seems reasonable toconclude that the inositol phosphates are quanti-tatively important in soil, although it is also clearthat their concentrations and contribution to thesoil organic phosphorus vary widely. This is illus-trated in Fig. 12.1, which shows solution 31PNMR spectra of the phosphorus extracted from

Inositol Phosphates in Soil 187

two US soils. One soil contains virtually all itsorganic phosphorus in the form of inositol phos-phates, while the other contains none. It shouldnot therefore be assumed that inositol phosphatesare abundant in all soils.

The majority of the inositol phosphates insoil are hexakisphosphates (i.e. with six phosphate

groups around the inositol ring). This is probablydue to a combination of factors, but principallythat most inputs to soil are as myo-inositol hexak-isphosphate from plants. The pentakisphosphatesare also abundant in some soils, altough lower-order esters (monophosphates to tetrakisphos-phates) are relatively rare (Omotoso and Wild,


−505

34567

Phosphate myo-Inositolhexakisphosphate

scyllo-Inositolhexakisphosphate

−505

Pyrophosphate

DNA

45

RNAmononucleotides

Phospholipiddegradationproducts

(a)

(b)

Fig. 12.1. The organic phosphorus composition of two soils from the USA with contrasting proportions ofinositol phosphates determined by extraction in sodium hydroxide and ethylenediaminetetraacetate (EDTA)and solution 31P nuclear magnetic resonance (NMR) spectroscopy. (a) The spectrum of a wetland soil froma nutrient-enriched site in the Florida Everglades that contained no detectable inositol phosphates. (FromTurner and Newman, 2005.) The soil was a Histosol containing 44% carbon and 0.16% phosphorus. (b) Thespectrum of an arable soil from Sussex County, Delaware, in which all detectable organic phosphorus in theextract was inositol hexakisphosphate. (From P. Murphy and B. Turner, 2003, unpublished data.) The soilwas an acidic sandy loam containing 0.7% carbon and 0.09% phosphorus. The zoomed inset spectra showthe phosphate monoester regions in detail. The main spectrum in (a) is plotted with 8 Hz line broadening,while all other spectra are plotted with 1 Hz line broadening to show fine resolution.

188B

.L.Turner

Table 12.1. Concentrations of inositol hexakisphosphate in surface soil from various parts of the world.

Inositol Proportion Number of Soil type and land Analytical hexakisphosphate of the total

Reference soils Location use Soil properties procedure (mg P/kg) organic P (%)a

Anderson 17 Scotland Range of textural Carbon 1.0–8.0% Hot 3 M NaOH, 56–460 24–58 (40)(1964) properties, mainly pH 4.7–7.5 ion-exchange

loams; land use Organic Pb 220– chromatographynot specified 920 mg P/kg

Caldwell 49 USA Range of virgin pH 4.9–7.8 Concentrated HCl, 2–62c 2–31 (12)and Black and cultivated Organic Pb 40– 0.5 M NaOH, (1958c) soils 446 mg P/kg ion-exchange

chromatographyIslam 20 Bangladesh Range of Ultisols, Organic matter Hot 3 M NaOH, 19–130d 21–58 (36)

and Mandal Entisols and 0.7–3.6% ion-exchange (1977) Inceptisols; land pH 4.7–8.1 chromatography

use not specified Organic Pb 63–261 mg P/kg

McKercher 18 Canada Range of soils; Nitrogen Hot 3 M NaOH, 20–71d 11–23 (17)and Anderson land use not 0.08–0.59%e ion-exchange (1968b) specified Organic Pb chromatography

100–475 mg P/kge

Thomas and 9 Canada Virgin loams Organic matter Concentrated HCl, 6–72c 2–10 (5)Lynch (1960) 2.4–13.6% 0.5 M NaOH,

pH 5.5–7.1 ion-exchange Organic Pb chromatography

148–710 mg P/kg

Inositol Phosphates in S

oil189

Turner (2006) 13 Madagascar Humid tropical Carbon NaOH–EDTA ND–33c ND–26Oxisols under rice 1.1–15.3% extraction, solution 31P

pH 4.6–5.8 NMR spectroscopyOrganic Pf

22–393 mg P/kgTurner et al. 29 England and Temperate lowland Carbon 2.9–8.0% NaOH–EDTA 26–189c 11–35 (22)

(2003b) Wales permanent pasture pH 4.4–6.8 extraction, solution 31Pwith high clay Organic Pf NMR spectroscopy(22–68%) 208–895 mg P/kg

Williams and 47 Australia Range of soils Nitrogen Hot 3 M NaOH, 1–356d <1–38 (16)Anderson and land use, 0.04–0.76% ion-exchange (1968) including cultivated, pH 5.0–9.0 chromatography

uncultivated and Organic Pg

pasture 6–1773 mg P/kg

ND = not detected; NMR = nuclear magnetic resonance.aValues in parentheses are the mean of all soils.bDetermined by the extraction method of Mehta et al. (1954).cValues are for myo-inositol hexakisphosphate only.dIncludes inositol pentakisphosphates.eValues estimated from figure.fDetermined by NaOH–EDTA extraction and solution 31P NMR spectroscopy.gDetermined by the extraction method of Saunders and Williams (1955).

1970a; Anderson, 1980). In fact, it remainsunclear whether the lower esters occur naturallyin soil or are an artefact of the strong chemicalsolutions necessary to extract inositol phosphatesfrom soil (Cosgrove, 1980).

The early literature contains many reportsof inositol phosphates in soil, but these must beassessed with care. Most studies prior to the1980s involved strong alkali extraction followedby column chromatography, which may haveinaccurately determined the soil inositol phos-phate content (Turner et al., 2002). Problemsincluded the incomplete recovery of inositolphosphate from anion-exchange columns(Anderson, 1964; Martin, 1970) and the inclu-sion of organic phosphorus compounds otherthan inositol phosphates in the separated frac-tions (Irving and Cosgrove, 1981). These prob-lems are exemplified by discrepancies observedin studies that compared the various methods.Dormaar (1967) measured greater inositol phos-phate concentrations in Canadian chernozemsusing the method of Caldwell and Black (1958a)compared with a procedure based on the meth-ods of Anderson (1956) and Cosgrove (1963). Indirect contrast, McKercher and Anderson(1968b) recorded less inositol phosphate inCanadian soils by the Caldwell and Black(1958a) method compared with the method ofAnderson (1964). The authors suggested that thismight have been caused by incomplete precipi-tation of iron-phytate when using the wideiron/inositol phosphate ratio of the Caldwell andBlack (1958a) procedure. However, Anderson(1964) had previously reported little differencebetween his method and that of Caldwell andBlack (1958a) for a range of British soils.

Recent studies employed a single-step alka-line extraction procedure with detection by solu-tion 31P NMR spectroscopy (Turner et al., 2003b,2005b; Turner and Richardson, 2004). Soilorganic phosphorus recovery in a solution con-taining sodium hydroxide and ethylenediaminetetraacetate (EDTA) is similar to the conventionalmethods used in the older literature (Bowman andMoir, 1993; Turner et al., 2005a), so presumablyresults from recent and earlier studies are broadlycomparable. Using solution 31P NMR spec-troscopy, it is currently possible to quantify myo-inositol hexakisphosphate by its four characteristicsignals that occur in a 1:2:2:1 ratio (Turner et al.,2003b), while scyllo-inositol hexakisphosphate is

identified by its single strong signal at the upfieldend of the phosphate monoester region (all sixphosphate groups are stereochemically identical)(Turner and Richardson, 2004). The signals canbe determined directly in extracts by spectraldeconvolution or following hypobromite oxidation(Fig. 12.2). The latter procedure destroys all phos-phate monoesters except the inositol phosphates(Wrenshall and Dyer, 1941) and convenientlyimproves the spectral resolution if EDTA isincluded in the NMR tube (Fig. 12.2; Turner andRichardson, 2004). Given that detection by NMRspectroscopy precludes the problems associatedwith column chromatography, NaOH–EDTAextraction and solution 31P NMR spectroscopyprovide a convenient and accurate alternative tothe conventional procedures for the determinationof inositol phosphates in soil. Further improve-ment is possible by using two-dimensional NMRspectroscopy (see Murthy, Chapter 2, thisvolume), although this has not yet been applied tothe complex matrices of soil extracts.

Factors Controlling the Amountsof Inositol Phosphates in Soil

Despite the large amount of published informa-tion on inositol phosphates in soil, there is still noclear understanding of the factors controllingtheir abundance. Indeed, the proportion of thesoil organic phosphorus as inositol phosphatescan vary appreciably, even in nearby soils formedunder similar environmental conditions andderived from the same parent material (Williamsand Anderson, 1968; Turner et al., 2003b).

In a meta-analysis of literature information,Harrison (1987) reported that ~90% of the vari-ation in the concentrations inositol phosphates insoils was explained by organic phosphorus, pHand organic carbon. This is relatively uninforma-tive, however, given that inositol phosphates typ-ically constitute most of the soil organicphosphorus. Studies of a large number of soilsunder similar climate and vegetation have indi-cated that inositol phosphates are not correlatedwith factors typically associated with organic mat-ter content, such as organic carbon, nitrogen, clayor microbial biomass. Rather, they are correlatedwith factors linked specifically to phosphate stabi-lization, such as the phosphate sorption capacity

190 B.L.Turner

and amorphous aluminium and iron (McKercherand Anderson, 1968b; Anderson et al., 1974;Turner et al., 2003b). This is unsurprising,because inositol hexakisphosphates are rapidlyand strongly sorbed to soil constituents and

can form insoluble precipitates with polyvalentcations (Jackman and Black, 1951; Anderson andArlidge, 1962). These abiotic processes accountfor the characteristic dynamics of inositol phos-phates in the environment and are reviewed in


(a) Phosphate

(b)

Unidentifiedinositol

phosphates

scyllo-Inositolhexakisphosphate

myo-Inositolhexakisphosphate

(c)

4567


Fig. 12.2. Identification of scyllo- and myo-inositol hexakisphosphates in a soil extract by hypobromiteoxidation and solution 31P nuclear magnetic resonance (NMR) spectroscopy. (From Turner andRichardson, 2004.) (a) The spectrum of an untreated sodium hydroxide/ethylenediaminetetraacetate(EDTA) extract of a Welsh pasture soil; (b) the same extract after hypobromite oxidation to destroy allphosphate monoesters other than higher-order inositol phosphates; (c) the same extract analysedwithout EDTA in the redissolved sample. Note the marked difference in resolution in the three spectra. Inspectrum (b) the strong signal at 6.2 ppm is inorganic phosphate, while all other signals are inositolphosphates. The soil was a clay loam containing 4.6% carbon and 0.11% phosphorus.

detail elsewhere in this volume (see Celi andBarberis, Chapter 13).

It therefore seems likely that inositolphosphates are stabilized in soil by differentmechanisms to those responsible for the stabiliza-tion of organic matter. This may explain in partthe discrepancies in nutrient stoichiometry observedin soil (McGill and Cole, 1981). However, at leastsome inositol phosphates are intimately associatedwith organic matter, because they have beendetected in high-molecular weight humic sub-stances isolated from soil (Moyer and Thomas,1970; Omotoso and Wild, 1970b). In addition, theproportions of phosphate monoesters in humicacids of lowland rice soil of the Philippines werepositively correlated with those of aromatic carbonand heterocyclic nitrogen, suggesting that inositolphosphates were contained within strongly humi-fied structures (Mahieu et al., 2002).

There is a decline in organic phosphoruswhen virgin soil is brought under cultivation(Tiessen et al., 1983), although inositol phosphatesappear to be depleted at a slower rate than otherorganic phosphates (Williams and Anderson, 1968;Condron et al., 1990). This selective mineralizationmeans that cultivated soils tend to contain a greaterproportion of their organic phosphorus as inositolphosphates (see Fig. 12.1). Some of the differencesmay be explained by sampling depth, becauseinositol phosphates accumulate at the surface ofundisturbed soil, but will be distributed throughoutthe plough layer in cultivated soil (Thomas andLynch, 1960; Dormaar, 1967; McKercher andAnderson, 1968b). They can, however, accumulatein the B-horizon of podzols (Williams andAnderson, 1968).

Several studies reported that temperate forestsoils contained more inositol phosphates thangrassland soils, both in terms of concentrationand as a proportion of the soil organic phosphorus(Caldwell and Black, 1958c; McKercher andAnderson, 1968b). Those studies rarely reportedthe type of forest, but this is likely to be impor-tant. For example, conversion from pasture topine forest in New Zealand caused a markeddecrease in soil organic phosphorus (Condronet al., 1996), which was subsequently linked to thedegradation of inositol phosphates (Chen et al.,2004).

It was recently reported that inositol phos-phates were absent from organic wetland soils ofthe Florida Everglades, USA (Turner and

Newman, 2005). As samples were analysed fromseveral locations with a range of chemical prop-erties, this suggests a major difference in thephosphorus cycle compared to mineral soils.Similar results were reported for constructedtreatment wetlands designed to sequester pollu-tant phosphorus from agricultural runoff (Turneret al., 2006). This raises important concernsabout the long-term stability of the organic phos-phorus sequestered in such systems, because itoccurs in relatively unstable phosphate diestersrather than recalcitrant inositol phosphates.

Although there are several possible explana-tions for the absence of inositol phosphates inthese wetland soils, the most likely is that anaero-bicity occurs close to the sediment surface formuch of the year. In acidic rice soil the hydroly-sis of inositol phosphates begins soon after sub-mergence (Islam and Ahmed, 1973), while inmarine sediments hydrolysis proceeds more rap-idly under anaerobic conditions than in parallelsamples incubated aerobically (Suzumura andKamatani, 1995). This may be due to the reduc-tion of ferric iron and the release of associatedinositol phosphates, which are then available forbiological attack. However, anaerobic reduction offerric iron–inositol hexakisphosphate complexeswas reported to form insoluble Fe4-phytate ratherthan release free inositol hexakisphosphate (DeGroot and Golterman, 1993). This could accountfor the observed decreases in inositol phosphatefollowing submergence of rice soils, but only if, asseems unlikely, the Fe4-phytate complex remainsinsoluble in the strong alkali used to extract inos-itol phosphates. This is also discussed elsewhere inthis volume (see Celi and Barbaris, Chapter 13,and McKelvie, Chapter 16).

Soil pH should exert a strong control on theaccumulation of inositol phosphates, becausesorption and metal precipitation are greaterunder acidic conditions ( Jackman and Black,1951; Anderson and Arlidge, 1962). In addition,phytases exhibit marked differences in behaviourdepending on pH – they are most active underacidic conditions, but are also inactivated bysorption to a greater extent in more acidic soil(see George et al., Chapter 14, this volume). Soilorganic phosphorus concentrations are oftennegatively correlated with soil pH (Harrison,1987), although this is less clear for the inositolphosphates. Caldwell and Black (1958c) reporteda significant negative correlation between soil pH

192 B.L.Turner


and the proportion of the soil organic phospho-rus as inositol phosphates, although other studies(e.g. Williams and Anderson, 1968) found nosuch relationship.

Inositol phosphates will readily form calciumprecipitates under alkaline conditions (see Celiand Barberis, Chapter 13, this volume), but thereis no clear evidence that the presence of carbon-ates influences inositol phosphate accumulation.Cosgrove (1966) reported the complete absence ofinositol phosphates in slightly alkaline calcareoussoils from Scotland, but other studies have meas-ured considerable quantities in calcareous soils(McKercher and Anderson, 1968b; Williams andAnderson, 1968). These discrepancies may beexplained by methodology, because Anderson(1964) reported a marked difference in the con-centration of inositol phosphates extracted from acalcareous soil by two different procedures.

There is also evidence for the relativesolubility of inositol phosphates in calcareous soils.In a study of a range of mainly cultivated calcare-ous soils from the semiarid western USA, concen-trations of phosphate monoesters determined bysolution 31P NMR spectroscopy of alkaline soilextracts were relatively small and not significantlycorrelated to carbonate content, although theywere correlated with other soil properties, includ-ing organic carbon, amorphous iron and alu-minium, pH and clay (Turner et al., 2003a).However, phytase-hydrolysable phosphorus assaysdemonstrated that a large proportion of theorganic phosphorus in bicarbonate extracts wasinositol hexakisphosphate, suggesting the relativesolubility and potential bioavailability of inositolphosphates in these calcareous soils. This was con-firmed by a recent study that demonstrated therapid decomposition of myo-inositol hexakisphos-phate in manure within weeks of application to acalcareous arable soil (see Leytem and Maguire,Chapter 10, this volume).

A key factor in the regulation of inositolphosphates in soil that has not been adequatelyaddressed is the role of nutrient status. Inositolphosphates are conventionally considered to berelatively unavailable to organisms due to theirstrong interaction with soil components (seeRichardson et al., Chapter 15, this volume), butmany soil microbes synthesize phytase and canutilize inositol phosphates in their environment(e.g. Richardson and Hadobas, 1997; Unno et al.,2005; see Hill and Richardson, Chapter 5, this

volume). This raises the possibility that inositolphosphates are used only when phosphorus isscarce relative to other nutrients. In other words,phosphorus limitation may drive the biologicaldegradation of recalcitrant inositol phosphates byfavouring organisms that can access them in soil.

Evidence in support of this hypothesis islimited, but a study of a large number of soilsunder similar vegetation and climate (temperatepermanent pasture) revealed that inositol phos-phate concentrations were greatest in soils with alow nitrogen/organic phosphorus ratios (Turneret al., 2003b, 2005b). In the case of myo-inositolhexakisphosphate, concentrations were alsogreater in soils containing more phosphate inreadily soluble form (as determined by bicarbon-ate extraction).

In agricultural soils, phosphate fertilizationwould be expected to decrease the biologicaldemand for phosphorus associated with soilorganic matter and lead to an accumulation ofinositol phosphate. This may explain in part theabundance of inositol phosphates in cultivatedsoils and their persistence compared with otherorganic phosphates such as nucleic acids andphospholipids during cultivation (Williams andAnderson, 1968; Condron et al., 1990). Analysisof soils of different ages from the Franz Josef gla-cial chronosequence in New Zealand revealed arapid increase in inositol phosphates (in terms ofboth concentration and proportion of the soilorganic phosphorus) during the first 1000 yearsof soil development, after which the concentrationsdeclined (Baker, 1977). As the early stages ofecosystem development are typically associatedwith nitrogen limitation (Walker and del Moral,2003), the abundance of inositol phosphatescould be linked to the plentiful availability ofphosphorus. The role of nutrient status thereforedeserves careful attention in future studies ofinositol phosphates in soil.

Phosphorylated InositolStereoisomers in Soil

Perhaps the most intriguing aspect of inositolphosphates in soil is the presence of stereoisomersother than myo-inositol that occur nowhere else innature. Nine stereoisomers of inositol exist (seeShears and Turner, Chapter 1, this volume),

although only four have been detected in soil inthe phosphorylated form (Fig. 12.3). Of these, onlymyo-inositol phosphates are prevalent in nature(Raboy, 2003). The remaining three stereoisomers(neo-inositol, D-chiro-inositol and scyllo-inositol),which differ from myo-inositol by the orientation ofa single hydroxyl group, occur widely in soil inhighly phosphorylated forms, yet are extremelyrare in biological tissue (discussed below).

Studies reporting quantitative estimates ofthe phosphorylated inositol stereoisomers in soilare summarized in Table 12.2. In agreementwith the results for total inositol phosphates, theconcentrations of phosphorylated stereoisomersand their contribution to the soil organic phos-phorus vary widely. The dominant phosphory-lated stereoisomer in soil is myo-inositol, as wouldbe expected from its widespread abundance inplants. The remaining isomers tend to occur inthe following order of abundance: scyllo-> D-chiro->neo-, although myo- and scyllo-inositol hexakispho-sphates together account for most of the soilinositol phosphate (McKercher and Anderson,1968a; Turner and Richardson, 2004). All threestereoisomers have been detected as lower estersin small concentrations (e.g. Halstead andAnderson, 1970). It should be noted, however,

that some studies did not detect scyllo-inositolphosphates, despite the presence of other phos-phorylated stereoisomers (Martin and Wicken,1966; L’Annunziata and Fuller, 1971), whileIrving and Cosgrove (1982) suggested that neo-inositol phosphates are not quantitatively deter-mined unless a hypobromite oxidation step isincluded in the analytical procedure. This meansthat neo-inositol phosphates may have beenunderestimated in parts of the older literature.

Early studies involving chromatographicseparation of inositol phosphates in soil extractsdocumented a ‘supposed isomer’ of myo-inositolhexakisphosphate that eluted at the end of thechromatogram (Smith and Clark, 1951; Caldwelland Black, 1958a) and was subsequently shownto be scyllo-inositol hexakisphosphate (Cosgrove,1962). Of the few studies in which quantitativevalues were obtained for a large number of soils,Caldwell and Black (1958c) reported scyllo-inosi-tol hexakisphosphate (although at the time ofpublication they did not identify it as such) toconstitute an average of 5.8% of the organicphosphorus in US soils, while concentrations intemperate pasture soils from England and Walesaveraged 9.7% of the soil organic phosphorus(Turner et al., 2005b). Concentrations of

194 B.L.Turner

HOOH

OH

OH

neo-Inositol

HO

HO

OHOH

2 axial groups

D-chiro-(+)-Inositol

HO

OH

OH OH

HO

No axial groups

scyllo-Inositol

1 axial group

myo-Inositol

HO

HO HO

HO

OH

OHHO OH

OH

HOHO

2 axial groups

Fig. 12.3. The four inositol stereoisomers that occur in phosphorylated forms in soil. Axial hydroxyls areshown in bold and hydrogen groups have been omitted. The four isomers differ only by the orientation ofa single hydroxyl group (circled). Compared with myo-inositol, D-chiro- and neo-inositols have an extraaxial group, whereas scyllo-inositol has none. This confers scyllo-inositol hexakisphosphate with aresistance to enzymatic attack that may account in part for its persistence in soils.


scyllo-inositol hexakisphosphate can exceed thoseof myo-inositol hexakisphosphate, although this isnot common (Table 12.2). Only a handful ofstudies have quantified D-chiro- and neo-inositolhexakisphosphate in soil, although Irving andCosgrove (1982) determined values in fourAustralian soils by gas chromatography. Ratiosof D-chiro- or neo-inositol phosphates to myo-inosi-tol phosphates are typically <0.2.

Quantitative data on the phosphorylatedinositol stereoisomers are fragmentary and insuf-ficient to draw firm conclusions on factors regu-lating their presence and abundance in soil.Concentrations of scyllo-inositol hexakisphosphatein a series of temperate pasture soils were corre-lated positively with soil organic phosphorus andmyo-inositol hexakisphosphate, although not withpH, organic carbon, amorphous iron and alu-minium or microbial biomass (Turner et al.,2005b). However, scyllo-inositol hexakisphosphatewas strongly correlated with the nitrogen/organic phosphorus ratio, suggesting that theabundance of this compound is regulated at leastin part by nutrient status. This was further sup-ported by the results of a 10-month pot experi-ment with six grassland soils from New Zealand,in which the growth of ryegrass (Lolium perenne L.)decreased scyllo-inositol hexakisphosphate inthree low-nutrient soils by 5–21%, but increasedit in three other high nutrient soils by 11–16%(Turner et al., 2005b). This indicates that organ-isms that are able to access inositol phosphatesmay be favoured when phosphorus is relativelyscarce, although further studies are clearlyrequired to assess this in greater detail.

Origins of Phosphorylated InositolStereoisomers

Despite their widespread occurrence in soils, thephosphorylated inositol stereoisomers other thanmyo have been detected rarely elsewhere innature and their origins in soil are unknown.Inositol phosphates are conventionally consid-ered to originate mainly from plants, but highlyphosphorylated stereoisomers other than myo-inositol have been detected only once in plant tis-sue. This suggests the importance of microbes inthe synthesis of these compounds, yet they havenever been detected in any soil organism.

Reports of the detection of inositolstereoisomers and their phosphorylated forms aresummarized in Table 12.3. myo-Inositol hexak-isphosphate is abundant in eukaryotes as both acellular component and in seeds (Raboy, 2003).In contrast, scyllo-inositol hexakisphosphate hasnever been detected in biological tissue, althougha phospholipid containing a scyllo-inositolmonophosphate occurs in barley aleurone (pro-tein stored as granules in the cells of plant seeds)(Kinnard et al., 1995; Narasimhan et al., 1997;Carstensen et al., 1999). The only known sourceof highly phosphorylated neo-inositol phosphatesis amoebae, including the freshwater carnivorousamoeba Amoeba discoide (Laird et al., 1976) and thehuman intestinal parasite Entamoeba histolytica(Martin et al., 2000). In the latter organism, neo-inositol occurs as both hexakisphosphate andpyrophosphate forms (i.e. with up to eight phos-phate groups), although the function of thesecompounds remains unclear.

The presence of D-chiro-inositol phosphatewas reported in needles of ponderosa pine (Pinusponderosa P. & C. Lawson) and leaves of velvetmesquite (Prosopis juliflora var. velutina (Sw.) DC.(Woot.) Sarg.) (L’Annunziata and Fuller, 1971),but it has never been detected in any other organ-ism. An additional phosphorylated stereoisomer,muco-inositol hexakisphosphate (Fig. 12.4), was alsodetected in the velvet mesquite leaves and is theonly report of this rare stereoisomer in any phos-phorylated form in nature. However, these resultswere subsequently questioned on analyticalgrounds, because the extracts probably containedconsiderable polysaccharide and nitrogenousmaterial that could have contained free inositols(Cosgrove, 1980). This is discussed in detail else-where in this volume (see L’Annunziata, Chapter 4).It would be appropriate to reassess similar samplesusing modern analytical techniques, given thepotential significance of the natural occurrence ofphosphorylated muco-inositol.

Stereoisomeric forms of inositol hexakispho-sphate were detected in aerobically digestedsewage sludge in similar ratios to those detectedin soils, although they constituted only around5% of the organic phosphorus (Cosgrove, 1973).As they were not present in raw sewage, it seemslikely that microbial activity was involved in theirsynthesis. However, only myo-inositol phosphateshave been reported in manures from a wide vari-ety of animals (including pigs, sheep, chickens,

Table 12.2. Studies reporting quantitative values for inositol hexakisphosphate stereoisomers other than myo in soils.

Proportion of Ratio of Number of Soil type and Analytical Isomers Concentration the total isomer to

Reference soils Location land use Properties procedure detected (mg P/kg soil) organic P (%) myo-

Caldwell 49 USA Range of pH 4.9–7.8 Concentrated HCl, scyllo - 1–39 1.6–21.1 0.17–1.18and Black virgin and Organic Pa 40– 0.5 M NaOH,(1958c) cultivated 446 mg P/kg ion-exchange

soils chromatographyCosgrove 3 Australia Two samples of Organic Pb 1 M NaOH, scyllo - 15–63 (scyllo) 2–3 0.20–0.25

(1963) alpine humus 770–2088 mg P/kg hypobromite (scyllo-and a basaltic oxidation, to myo-+soil ion-exchange D-chiro-)

chromatographyIrving and 4 Australia Not reported Not reported Hot 3 M NaOH, scyllo - 4.2–33.1c – 0.30–0.41

Cosgrove hypobromite D-chiro- 1.2–13.1c 0.09–0.12(1982) oxidation, gas neo - 1.2–10.0c 0.09–0.11

chromatographyMartin and 5 New Zealand Range of Carbon 6.6–12.3% 0.3 M KOH, D-chiro- 4.0–42c 0.37–0.56

Wicken surface soils Organic Pd ion-exchange (1966) 440–1360 mg P/kg chromatography

McKercher 8 Canada and Range of arable, Carbon Hot 3 M NaOH, scyllo - 9.0–87.4c 3.4–16.2c 0.22–0.90and Scotland grassland and 1.8–6.6% ion-exchange D-chiro- <5 (scyllo-Anderson forest soils pH 4.9–7.8 chromatography neo - <1 to myo- +(1968a) Organic Pa D-chiro)

200–920 mg P/kg

Omotoso 7 England and Grassland and Carbon 1.5–13.1% 1 M NaOH, scyllo- ND–5.1e 0–0.32and Wild Nigeria forest soils pH 3.7–7.2 hypobromite D-chiro- 1.1–2.9e 0.07–0.18(1970a) Organic Pa oxidation, neo - ND–0.4e 0–0.02

17–175 mg P/kg ion-exchange chromatography

Thomas 9 Canada Virgin loams Organic matter Concentrated HCl, scyllo - 2.8–83.4 <1–12 0.45–1.83and Lynch 2.4–13.6% 0.5 M NaOH, (1960) pH 5.5–7.1 ion-exchange

Organic Pb chromatography148–710 mg P/kg

Turner (2006) 13 Madagascar Humid tropical Carbon 1.1–15.3% NaOH–EDTA scyllo - ND–44 ND–11 0.33–0.67Oxisols under pH 4.6–5.8 extraction, rice Organic Pf solution31P NMR

22–393 mg P/kg spectroscopyTurner et al. 29 England Temperate Carbon 2.9–8.0% NaOH–EDTA scyllo- 11–130 4.4–14.5 0.29–0.79

(2005b) and Wales permanent pH 4.4–6.8 extraction, lowland pasture Organic Pf solution 31P NMR with high clay 208–895 mg P/kg spectroscopy(22–68%)

ND = not detected; NMR = nuclear magnetic resonance.aDetermined by the extraction method of Mehta et al. (1954).bDetermined by the extraction method of Saunders and Williams (1955).cIncludes inositol pentakisphosphates.dDetermined by the ignition method of Saunders and Williams (1955).eValues are for all esters of the stereoisomer (i.e. from monophosphates to hexakisphosphates).fDetermined by NaOH–EDTA extraction and solution 31P NMR spectroscopy.

198 B.L.Turner

Table 12.3. The occurrence of phosphorylated and free inositol stereoisomers in nature.

Isomer Axial groups Occurrence in phosphorylated form Occurrence of the free inositol

scyllo- 0 Common in soils as the Widespread in plants, Inositol hexakisphosphate and lower esters including chrysanthemums

(Cosgrove, 1962); also detected in (Ichimura et al., 2000) and aerobic sewage sludge (Cosgrove, Proteaceae (Bieleski and 1973); a phospholipid containing scyllo- Briggs, 2005); also detected inositol monophosphate occurs in the ciliate Tetrahymena in barley aluerone cells (Kinnard vorax (Kersting et al., 2003),et al., 1995) the red algae Porphyra

umbilicalis and a number of other organisms (Posternak, 1965)

myo-Inositol 1 Abundant as the hexakisphosphate Widespread in biological and lower esters in a range of tissue (Morré et al., 1990;organisms (Raboy, 2003) and soils Loewus and Murthy, 2000)(Turner et al., 2002)

D-chiro- 2 Abundant as the hexakisphosphate Occurs in plants, often in Inositol in soil, with smaller amounts of the methylated form as pinitol

lower esters (Cosgrove, 1969); also (Morré et al., 1990); also detected in aerobic sewage sludge detected in trace amounts (Cosgrove, 1973), pine needles in the ciliate Tetrahymena and velvet mesquite leaves vorax (Kersting et al., 2003)(L’Annunziata and Fuller, 1971)

L-chiro- 2 Never detected in phosphorylated form Occurs in plants such as Inositol seagrass (Drew, 1983),

chrysanthemums (Ichimura et al., 2000) and Proteaceae (Bieleski and Briggs, 2005), often in methylated form

neo -Inositol 2 Occurs in soils as the hexakisphosphate Occurs in mammalian tissue and occasionally lower esters (Sherman et al., 1971), but (Cosgrove and Tate, 1963); also rare in plants (Mukherjee detected in aerobic sewage sludge and Axt, 1984); detected in (Cosgrove, 1973); highly trace amounts in the phosphorylated forms occur in protozoan Tetrahymena amoebae living in human intestines vorax (Kersting et al., 2003)(Martin et al., 2000) and fresh water (Laird et al., 1976); also present as the monophosphate in mammalian brain tissue (Sherman et al., 1971)

epi -Inositol 2 None No natural sourcemuco -Inositol 3 Detected once in velvet mesquite Occurs in plants such as

leaves (L’Annunziata and Fuller, 1971) gymnosperms, a few angiosperms (Dittrich et al.,1971; Dittrich and Kandler, 1972) and seagrass (Drew, 1983), often in methylated form

allo -Inositol 3 None No natural sourcecis -Inositol 3 None No natural source


turkeys, dairy and beef cattle) using solution 31PNMR spectroscopy, despite a detection limit ofaround 1 mg P/kg dry manure (Leytem et al.,2004; Maguire et al., 2004; Turner, 2004).

Chemical synthesis is unlikely to be involvedin the formation of the phosphorylated stereoiso-mers in the environment (Cosgrove, 1980). It canoccur following extended heating in strong acid oralkali (Cosgrove, 1975) or through simple reactionssuch as the epimerization of myo-inositol pentak-isphosphate to scyllo- and D-chiro-inositol pentak-isphosphate by hypobromite oxidation followed bysodium borohydride reduction (Cosgrove, 1972).However, such reactions would clearly not occurin soil.

Unphosphorylated inositol stereoisomers, bycomparison to the phosphorylated forms, are rel-atively widespread in nature (Table 12.3). Fiveare common in plant tissue (myo-, D-chiro-, L-chiro,scyllo- and muco-), often in methylated forms suchas pinitol (3-O-methyl D-chiro-inositol) that play arole in stress tolerance (Loewus and Murthy,2000). A sixth stereoisomer, neo-inositol, occurs inmammalian tissue (Sherman et al., 1971), but hasbeen reported only once in a plant (Mukherjeeand Axt, 1984). The remaining three inositols (cis-,epi- and allo-) have no natural source.

Only myo-inositol appears to be synthesizeddirectly in plants, with other stereoisomersformed by epimerization reactions (Loewus andMurthy, 2000). Such reactions are similar to theconversion of myo-inositol to scyllo-inositol inbovine brain, which involves the enzymaticepimerization of the C-2 carbon of myo-inositol(Hipps et al., 1973). Synthesis can also occur bycyclization of an appropriate sugar phosphate, asin the formation of neo-inositol 1-phosphate in ratbrain via cyclization of mannose 6-phosphate byL-myo-inositol 1-synthase (Sherman et al., 1971).

Several laboratory studies have detected theformation of inositol phosphates in ‘synthetic’soils incubated with sugar and nutrients, whichsuggests a microbial source for the phosphory-lated stereoisomers (Table 12.4). Smith and Clark(1951) demonstrated that radiolabelled phosphateadded to soil was incorporated into inositol hexa-kisphosphate after incubation for 30 days withdextrose and ammonium nitrate. Examination ofthe chromatograms suggests that both myo- andscyllo-inositol hexakisphosphates were present inan approximate 2:1 ratio. This provides unequiv-ocal evidence for the biosynthesis of inositol phos-phates by soil organisms, although the specificprocess leading to incorporation of the labelledphosphate into inositol hexakisphosphates wasnot resolved (see below).

Direct synthesis of inositol phosphates by soilorganisms was also demonstrated by incubating‘synthetic’ soils, or inositol phosphate-free soil,with sugar, inorganic nutrients and a water extractof soil to inoculate the samples with soil microbes(Caldwell and Black, 1958b). Both myo- and scyllo-inositol hexakisphosphates were subsequentlydetected, with the scyllo form being more abun-dant in every sample. Cosgrove (1964) was subse-quently able to detect only myo-inositolhexakisphosphate in a similar, albeit shorter,experiment, although it is possible that other phos-phorylated stereoisomers were present in unde-tectable concentrations. Of potential significanceis that inositol phosphates were not detected insamples maintained at pH 6, whereas ~3 mgP/kg were detected in two more acidic samples.

Important insight into the origin of thephosphorylated stereoisomers was provided byexperiments in soil using 14C-labelled compounds(Table 12.4). These demonstrated unequivocallythat myo-inositol can be epimerized to D-chiro-inositol, and that myo-inositol can be phosphory-lated to the hexakisphosphate (L’Annunziata andGonzalez, 1977; L’Annunziata et al., 1977). Thephosphorylation of D-chiro-inositol was notdetected, but the study nevertheless providedstrong evidence for a microbial source of thephosphorylated stereoisomers in soil.

The relative abundance of free inositol iso-mers in plants raises the possibility that theycould be a source of the free isomers in soil.However, the fact that two isomers common inplant tissue (L-chiro- and muco-inositol) do notoccur in phosphorylated forms in soil suggests

PO

PO

PO

OP

OP

OP

Fig. 12.4. The structure of muco-inositolhexakisphosphate, which was reported once inplant tissue but never in soil. Phosphate groupsare denoted by ‘OP’.

200B

.L.Turner

Table 12.4. Studies showing the synthesis of inositol phosphates by soil organisms in laboratory incubations.

Reference Soil properties Nutrients added Incubation conditions Extraction and analysis Inositol phosphates detected

Smith and A loam under prairie Dextrose, ammonium Field capacity, room 0.5 M NaOH, hypobromite Radiolabelled myo- and scyllo-Clark (1951) (pH 6.7, organic P nitrate and 32P temperature, 30 days oxidation, ion-exchange inositol hexakisphosphates

144 mg P/kg) and a chromatography (~2:1 ratio) plus some silt loam under forest pentakisphosphates(pH 5.3, organic P 289 mg P/kg)

Caldwell and Various samples of soil Sucrose, inorganic 10 months at 30°C, Concentrated HCl, 0.5 M myo-Inositol hexakisphosphate Black (1958b) C-horizons (calcareous nutrients (also or 3 months at 28°C NaOH, ion-exchange (0–1.16 mg P/kg) and scyllo -

and acidic), sand and soluble starch in chromatography inositol hexakisphosphate sand–clay mixtures, first experiment) and (0.12–2.14 mg P/kg); scyllo >with pH between soil–water extract myo in all samples5.1 and 7.3

Cosgrove (1964) Kaolinite and Sucrose, inorganic 5 months, 25°C 1 M NaOH, hypobromite 3.0 mg P/kg myo-inositolsand (1:4 ratio) nutrients and soil– oxidation, ion-exchange hexakisphosphate in the two maintained at three water extract chromatography most acidic samples, none inpH values (4, 5 and 6) the pH 6 sample

L’Annunziata A-horizon of an 14C-myo -Inositol Field capacity, Hot 3 M NaOH, ion- 14C-myo-Inositolet al. (1977) uncultivated Andisol 12 days, 36.5°C exchange chromatography hexakisphosphate, 14C-D-chiro-

under forest (pH 5.8, inositolorganic matter 11%, clay 18%)


that an alternative source is more likely. Of pos-sible significance, therefore, is that the freshwater ciliate Tetrahymena vorax contains the samefour inositol stereoisomers in similar proportionsto those present in phosphorylated forms in soil(Kersting et al., 2003). Only the free inositolswere detected, although a series of higher-ordermyo-inositol phosphates were identified inParamecium tetraurelia, another fresh water proto-zoan (Freund et al., 1992). As protozoa are abun-dant in soils, they may therefore be an importantsource of the inositol stereoisomers in free orphosphorylated forms.

Despite the evidence for a microbial sourceof the phosphorylated stereoisomers, no soilorganisms have so far been found to containthem. There is no known prokaryotic source ofinositol phosphates. Of the eukaryotic microor-ganisms, Cosgrove (1964) could not detect inos-itol phosphates in the yeast Saccharomycescarlsbergensis or in 12 fungi isolated from soilsrich in phytate. However, S. cerevisiae (Baker’syeast) contains abundant myo-inositol phos-phates, including highly phosphorylated inositolpyrophosphates that appear to phosphorylateproteins (Saiardi et al., 2004). The latter processis selective for eukaryotic proteins, as it was notobserved in bacterial extracts.

The Potential Function ofPhosphorylated Inositol

Stereoisomers in Soil

Why do soil organisms synthesize phosphorylatedinositol stereoisomers such as scyllo-inositol hexa-kisphosphate when nature relies almost exclu-sively on the myo stereoisomer? What are theevolutionary benefits of stereoisomeric inositolphosphates that have favoured their synthesis insoils? And which organisms synthesize them?These are perhaps the most intriguing questionsregarding inositol phosphates in soil, but weseem far from answering them.

The obvious functional difference betweenmyo-inositol hexakisphosphate and the other phos-phorylated stereoisomers that occur in soil is therelative resistance of the latter to enzymaticattack. In particular, scyllo-inositol hexakisphos-phate, with no axial groups, seems to be mostresistant to phytase activity and is also the most

abundant phosphorylated stereoisomer other thanmyo-inositol found in soil. For example, Greaveset al. (1967) reported that phytase isolated fromAerobacter aerogenes, a Gram-negative facultativeanaerobic bacteria, had no activity towards scyllo-inositol hexakisphosphate. Cosgrove (1970) subse-quently found that the ‘SB2’ phytase from a soilPseudomonas sp. was active towards scyllo-inositolhexakisphosphate, but that the rate of hydrolysiswas the slowest of the four isomers tested, beingin the order of myo-> neo-> D-chiro-> scyllo-. Otherphytases, such as β-propeller phytase and purpleacid phytase (see Mullaney and Ullah, Chapter 7,this volume), have not yet been tested againstphosphorylated stereoisomers other than myo-inositol hexakisphosphate.

The resistance of scyllo-inositol hexakisphos-phate to hydrolytic attack raises the possibilitythat it might be synthesized by certain organismsto protect phosphorus from uptake by nearbycompeting organisms. Such a strategy for con-serving phosphorus might be expected to occur inenvironments where phosphorus is scarce, andcould conceivably arise among soil microbes or aspart of the complex competition between plantsand microbes for soil nutrients (Kaye and Hart,1997). There is currently no evidence to supportthis hypothesis, but it warrants investigation.

A further possibility is that stereoisomericinositol phosphates have a non-nutritional func-tion in soils. These highly reactive, but biologicallyrecalcitrant, compounds might, for example, playa role in soil structure (Anderson, 1980) by stabi-lizing clay–metal–humic complexes in an analo-gous way to their presence in the core of humanRNA-editing enzymes (Macbeth et al., 2005).Alternatively, their capacity to form insoluble pre-cipitates with metals might mean that they aresynthesized to ameliorate metal toxicity. Forexample, the arsenic hyperaccumulating brakefern Pteris vittata secretes inositol hexakisphosphatefrom its roots (Tu et al., 2004), although it is notknown which stereoisomers are involved.

Conclusions and Research Priorities

Soils contain large amounts of inositol phosphates,some of which occur nowhere else in nature, yetour understanding of their origin and function insoils is extremely limited. This is unsatisfactory

given the importance of soil phosphorus in theregulation of ecosystems, nourishment of cropsand pollution of water bodies. Future researchshould focus on quantifying the concentrations ofthe various stereoisomers of inositol hexakisphos-phate in soil from a wide range of environmentsand on investigating their biochemical origin andfunction. In particular, efforts should be directedtowards the following key topics:

● Improvement in analytical procedures for quantifica-tion of inositol phosphates in soils. Recentadvances in the analysis of soil inositol phos-phates by solution 31P NMR spectroscopymean that the quantification of myo- andscyllo-inositol hexakisphosphate is nowstraightforward. This should reinvigorateresearch on these compounds, although con-tinued improvements are of key importance.In particular, the ability to rapidly quantifyall four stereoisomeric inositol phosphatesthat occur in soils is desirable. The use oftwo-dimensional NMR spectroscopy or massspectrometry may facilitate this (see Murthy,Chapter 2, and Cooper et al., Chapter 3, thisvolume).

● Assessment of the forms and concentrations of inos-itol phosphates in soil from a wide range of envi-ronments. Inositol phosphates are abundantin most mineral soils, but all or none of thesoil organic phosphorus can be in this form.More information is therefore required,especially on the phosphorylated stereoiso-mers, for soil from a variety of environ-ments and ecosystems. Most data are fromtemperate agroecosystems that are relativelyfertile compared to soil under natural vege-tation. Little is available for the other majorbiomes, but this is likely to provide impor-

tant information on the dynamics and func-tion of the inositol phosphate stereoisomersin soil.

● Investigation of the dynamics of inositol phosphatesin soil. Inositol phosphates are consideredto be recalcitrant in soil, yet recent evi-dence suggests that they are a potentiallyimportant source of phosphorus to organ-isms. In particular, microbes with thecapacity to utilize inositol phosphates seemto be widespread in the environment andmay be important in making inositolphosphates available to plants. More infor-mation is now required on the rates of syn-thesis and decomposition of inositolphosphates in soil, particularly for the phos-phorylated stereoisomers. This will almostcertainly involve the use of isotopes (seeL’Annunziata, Chapter 4, this volume).

● Determination of the origins of the phosphorylatedinositol stereoisomers in soil and the organismsinvolved in their synthesis. More than threedecades ago, Cosgrove (1972) wrote that‘[f]inal proof of a microbial origin for theisomers awaits the isolation of soil organismscapable of their biosynthesis’. However, noorganism that synthesizes inositol hexak-isphosphate stereoisomers other than myo

has yet been isolated. The rapid advancesbeing made in DNA-based communityanalysis should facilitate the investigation ofa microbial origin for the stereoisomers,although the possibility remains that othertypes of organisms, including plants, are alsoimportant sources. Confirmation of abiosynthesis pathway for the phosphorylatedinositol stereoisomers would be a majoradvance in our understanding of these enig-matic compounds.

202 B.L.Turner

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13 Abiotic Reactions of InositolPhosphates in Soil

Luisella Celi and Elisabetta BarberisUniversity of Turin, DIVAPRA Chimica Agraria, via Leonardo da Vinci 44,

Grugliasco, 10095 Torino, Italy

Although biotic reactions are the main processesgoverning the transformation of organic matterin biologically active soils and sediments, abioticreactions can indirectly affect the fate of organiccompounds and control their persistence in theenvironment. The accumulation of organic phos-phorus in soil, often reaching 80% or more of thetotal phosphorus (Anderson, 1980), is attributedto a series of abiotic processes that hamper thebiodegradation of certain compounds (Anderson,1980; Stewart and Tiessen, 1987; Condron et al.,1990). These processes are related mainly to thehigh affinity of organic phosphorus for soil min-eral colloids, which have a large surface area anda large capacity to retain anions (Tiessen et al.,1983; Guzel and Ibrikci, 1994). Complexationand precipitation with polyvalent cations mayalso enhance the retention of organic phosphorusin the colloidal phase. This explains why organicphosphorus generally accumulates in the finestsoil fractions (Fig. 13.1), with the clay fractionoften containing more organic phosphorus thansilt (Gburek et al., 2005).

Sorption and precipitation with soil cationslimit in particular the degradation of phosphatemonoesters compared with phosphate diestersand other organic phosphorus compounds. Thelatter are less strongly stabilized and more likelyto be found in soil solution, where they are read-ily degraded by biological processes. This leads tothe accumulation of phosphate monoesters in

soils (Table 13.1), although the distribution oforganic phosphorus compounds in the livingorganisms from where they originate is quite dif-ferent (Magid et al., 1996).

In most soils the inositol phosphates are themost abundant group of phosphate monoesters;they occur in various degrees of phosphorylation(from inositol hexakisphosphate to inositolmonophosphate) and up to four stereoisomericconfigurations (myo-, scyllo-, neo-, D-chiro-) (Cosgrove,1980; Harrison, 1987; Turner et al., 2002). Theamounts and forms of inositol phosphates in soilsare reviewed elsewhere in this volume (see Turner,Chapter 12). This chapter examines the main abiotic processes involved in the stabilization ofinositol phosphates in soil. These include adsorp-tion and desorption from soil components, com-plexation reactions and precipitation withpolyvalent cations. The influence of soil solutionchemistry on these reactions is also discussed, aswell as the effects of inositol phosphates on soil surface properties.

Adsorption of Inositol Phosphates in Soil

Phosphate adsorption is one of the most widelystudied reactions in soil, whereas adsorption oforganic phosphorus compounds has received

208 L. Celi and E. Barberis

much less attention. From a theoretical point ofview, the term sorption is used instead of adsorp-tion, because it covers any process that removesreactant from the solution, including adsorptionand precipitation (Barrow, 1985, 1993).

The extent of adsorption of inositol phos-phate is controlled by its concentration in solu-tion, whereas the extent of precipitation isdetermined by the solubility product of the leastsoluble inositol phosphate compound. This inturn controls the anion concentration in solution,so the extent of inositol phosphate adsorption isalso linked to the nature and concentration ofpolycations in the system that can form insolublesalts. Moreover, the nature of the inositol phos-phates (i.e. the number of phosphate groups andthe stereochemical configuration) can furtheraffect both the arrangement of adsorbates on theactive surface and the formation of precipitates.

Adsorption is also governed by the charac-teristics of the adsorbate. Soil properties such aspH, mineral composition and texture (Fig. 13.2)can strongly affect the rate and extent of inositol

phosphate adsorption (Anderson et al., 1974;McKercher and Anderson, 1989; Leytem et al.,2002). The greater affinity of some soil compo-nents towards these organic compounds is relatedto their specific surface and porosity, degree ofcrystallinity and surface charge, and is stronglycontrolled by the pH of the system.

The role of iron and aluminium oxides

In acid soils the sorption of inositol phosphates isreported to be dependent on the contentsof amorphous iron and aluminium oxides(Anderson et al., 1974; Harrison, 1987; Pant et al.,1994). Among the different iron oxides, theamorphous types such as ferrihydrite alone, or inassociation with kaolinite, show a greater capac-ity to retain myo-inositol hexakisphosphate (Table13.2) compared to the more crystalline goethiteor haematite (Ognalaga et al., 1994; Celi et al.,1999, 2003). The presence of aluminium in the

00.0

2.0

4.0

6.0

8.0

10 20 30Clay (%)

40 50

Org

anic

P e

nric

hmen

t rat

io

Fig. 13.1. Relationship between the organic phosphorus enrichment ratio and the content of clay indifferent types of soil. (From Gburek et al., 2005.)

Table 13.1. Distribution of organic phosphorus fractions (as % of total phosphorus) in soils and growingorganisms. (From Magid et al., 1996.)

Escherichia coli Fungi Nicotiana Soils

Nucleic acids 65 58 52 2Phospholipids 15 20 23 5Phosphate Monoesters 20 22 25 50

Abiotic Reactions in Soil 209

goethite structure does not affect the extent ofadsorption (Cannoni et al., 2004). With theexception of ferrihydrite, the iron oxides show ahigher affinity for inositol phosphates than forother organic and inorganic phosphates, asdeduced by Langmuir K values (Table 13.2).

Adsorption of inositol phosphates occursthrough their phosphate groups, which react withiron oxide in the same way as the free phosphateion via a ligand exchange with the H2O and–OHgroups of the surfaces (Parfitt et al., 1976;Goldberg and Sposito, 1985). Evidence of thishas been obtained by a combination of quantita-tive, electrochemical and spectroscopic studies.The different sorption ratios observed betweenmyo-inositol hexakisphosphate and phosphate onthe various oxides indicate the involvement of avariable number of phosphate groups in the for-mation of the bonding and a different arrange-ment of the molecule depending on thecharacteristics of the surface (Celi et al., 1999,2001a, 2003). The number of active sites, the dis-tances between contiguous hydroxyls and theroughness of the mineral surface are the mainfactors affecting the phosphorus–mineral com-plex configuration. The phosphate groups thatdo not react with the surface remain free andmake the surface highly negative. The electricalpotential of the new surfaces formed by the phos-

phorus–iron oxide complex is related to theextent of adsorption and to the number of phos-phate groups that remain free after adsorption(Celi et al., 1999). Fourier-transform infraredspectroscopy shows changes in the P=O andP–O bands of myo-inositol hexakisphosphate afteradsorption on iron oxides (Fig. 13.3), due to theformation of Fe–O–P bonds (Celi et al., 1999).The higher electron-donor effect of the O–Fecompared with the –OH group caused a repul-sion of electrons to oxygen to form Pδ+–Oδ−,lowering the bond energy of P=Ο and increasingthat of P–O (Socrates, 1980).

From these results, Ognalaga et al. (1994) andCeli et al. (1999) suggested that adsorption of myo-inositol hexakisphosphate on goethite occurredthrough four of the six phosphate groups, with theremaining two groups being free. On ferrihydriteonly two phosphate groups were involved, dueprobably to the roughness of the oxide preventingthe optimal arrangement on the surface (Celi et al.,2003). Two phosphate groups per molecule of inos-itol phosphate are also bound to the surface ofhaematite, due in this case to an unfavourable dis-tance between the –OH sites for an optimalarrangement of the molecule on the less reactivesurface (L. Celi et al., unpublished data).

If iron oxides are associated with other soilcomponents, such as kaolinite, the surface area

00

400

800

Sor

bed

P (

µg/g

)

1200

1600

2000

20 40

Equilibrium P (mg P/m)

60 80 100

myo-IP6 sandymyo-IP6 sandy claymyo-IP6 sandy loammyo-IP6 clayPi sandyPi sandy clayPi sandy loamPi clay

Fig. 13.2. Isotherms of adsorption of myo-inositol hexakisphosphate (myo-IP6) and inorganic phosphate(Pi) in different types of soil. (Adapted from McKercher and Anderson, 1989; Leytem et al., 2002.)


and porosity are closer to that of the phyllosili-cate, while the electrical charge is governed bythe iron oxide (Celi et al., 2001b). Adsorption ofinositol phosphate is similar to that of ferrihydritein quantitative terms, despite these different min-eralogical characteristics, probably due to a dif-ferent arrangement of the molecule. On themixed system inositol hexaphosphate is adsorbedprobably through only one of its six phosphategroups (Celi et al., 2003).

The affinity of inositol phosphates for oxidesseems to be related to the number of phosphategroups involved in the bonding: the higher thenumber of phosphate groups bound to the sur-face, the lower the affinity (see Langmuir con-stants, Table 13.2) as a result of the higher energynecessary to form the multiple bonding and toovercome the conformational hindrance deter-mined by the organic moiety (Celi et al., 1999,2003). A relatively strong correlation (r2 = 0.877;n = 5) was found between the Langmuir K valuesfor goethite, kaolinite, illite, ferrihydrite andmixed ferrihydrite–kaolinite systems, and the inos-itol hexakisphosphate/inorganic phosphate maxi-mum molar ratio (Fig. 13.4; Celi et al., 2003).

In acidic soils, aluminium oxides can alsopromote the retention of organic phosphorus(Anderson et al., 1974), although to a lower extent

than iron oxides. The capacity of amorphous alu-minium oxides to remove myo-inositol hexak-isphosphate from solution (Shang et al., 1990,1992) was much greater than that of bohemite(Anderson et al., 1974) and bayerite (Cannoniet al., 2004), due to the larger surface area.Anderson and Arlidge (1962) observed a limitedcapacity of gibbsite to adsorb inositol phosphates.myo-Inositol hexakisphosphate has a higher chem-ical affinity to the aluminium oxide surface com-pared to myo-inositol monophosphate, confirmingthat the interaction is regulated by the functional-ity of the phosphate groups (Shang et al., 1990,1992). For instance, three phosphate groups permolecule of myo-inositol hexakisphosphate wereinvolved in bonding to bayerite. This results in ahigh thermodynamic stability of myo-inositolhexakisphosphate–aluminium oxide complexesand a low activation energy with a higher rateconstant. As for inorganic phosphate, adsorptioninitially proceeds due to the high concentrationof adsorbate in the surrounding particle,although studies of this topic are limited. As sur-face coverage proceeds, the adsorbed myo-inositolhexakisphosphate can impede the approach ofother molecules to the surface by imposing nega-tive electrical and steric effects (Shang et al.,1990). As for sorption to iron oxides, desorption

Table 13.2. Langmuir coefficients of adsorption isotherms of myo-inositol hexakisphosphate (myo-IP6)and phosphate on iron and aluminium phyllosilicates, and ferrihydrite–kaolinite systems (Fh–KGa2) atpH 4.5 in 0.01 M KCl. (From Celi et al., 1999, 2003; Cannoni et al., 2004.) Xmax is the maximum amountadsorbed on the minerals while K is the Langmuir constant and indicates the affinity of the adsorbate forthe surface.

xmax (µmol/m2)a K (l/mol) r 2 (n = 10)

Goethite myo-IP6 0.64 (3.8) 8.0�103 0.978Phosphate 2.4 5.6�103 0.996

Ferrihydrite myo-IP6 2.12 (12.7) 2.4�105 0.994Phosphate 4.57 3.2�105 0.996

Haematite myo-IP6 0.40 (2.4) 6.2�104 0.993Phosphate 0.82 4.0�104 0.994

Bayerite myo-IP6 0.40 (2.4) 1.3�105 0.993Phosphate 1.3 7.7�104 0.994

Illite myo-IP6 0.38 (2.3) 1.0�105 0.993Phosphate 1.0 7.5�103 0.994

Kaolinite myo-IP6 0.27 (1.6) 1.0�105 0.999Phosphate 0.79 1.6�103 0.923

(Fh–KGa2) myo-IP6 2.24 (13.4) 2.2�106 0.993Phosphate 2.96 9.1�104 0.991

aThe values in parentheses indicate the amount of adsorbed myo-inositol hexakisphosphate expressed as moles ofphosphorus.


from aluminium precipitates is limited, and nei-ther plants nor microorganisms are able torelease the phosphate groups that remain freeafter adsorption (Shang et al., 1996), indicatingthat phosphorus bioavailability and biodegrad-ability are strongly limited by the stability ofphosphate surface complexes.

The role of calcium carbonate,clays and organic matter

In neutral and basic soils the sorption of inositolphosphates is governed by calcite, clays andorganic matter (McKercher and Anderson, 1989).Calcareous soils are those with the ability toimmobilize large amounts of phosphate, due tothe importance of calcium carbonate on phos-

phate chemistry (Ryan et al., 1984). Calcite canretain myo-inositol hexakisphosphate from solutionin amounts that largely exceed the maximumcoordination capacity of the surface (Celi et al.,2000). This occurs because, in addition to adsorp-tion on the reactive surface, inositol phosphatecan complex the calcium ion in equilibrium withthe mineral and form two soluble calcium phytatespecies, Ca1-phytate and Ca2-phytate, even at avery low concentration of phosphorus, whereasthe Ca3–phytate complex precipitates at any pH(Table 13.3; Graf, 1983). Calcium complexationcan favour the further dissolution of calcite andhence enhance precipitation of Ca3-phytate. Thiscan lead to the accumulation of organic phospho-rus in soil dependent on CaCO3-specific surfacearea, which is related to the particle size ratherthan to the total amount of CaCO3 (Holford andMattingly, 1975; Amer et al., 1985).

4000 3000

3174

Abs

orba

nce

minus GtGt−myo-IP6

myo -IP6

1646

1642 1385

1223

292733723397

1038 629

639

640794

889

794890

498798

9911078

1125

829433

792897

3123

3119Goethite

Gt−myo-IP6

2000

Wave number (1/cm)

1500 1000 400

Fig. 13.3. Fourier-transform infrared spectra of goethite (Gt), myo-inositol hexakisphosphate (myo-IP6),the complex formed between myo-inositol hexakisphosphate and goethite (Gt–myo-IP6) and thedifference spectrum obtained by subtracting the spectrum of Gt from that of the Gt–myo-IP6 complex(Gt–myo-IP6 minus Gt). (From Celi et al., 1999.)


Clay minerals can also be responsible for theretention of inositol phosphates in neutral orbasic soils, although their sorption capacity isless with respect to iron and aluminium oxides.Montmorillonite, and to a much smaller extentbentonite, illite and kaolinite, can adsorb myo-inositol hexakisphosphate (Goring andBartholomew, 1950; Celi et al., 1999). At pH 4.5,illite adsorbs more than kaolinite (0.38 and 0.27µmol/m2, respectively), while both phyllosilicatesshow a higher affinity (>KL) than for phosphate,with a better fitting of the Langmuir model evenat high adsorbate concentrations (Celi et al.,1999). This could be due to the fact that the

steric hindrance of myo-inositol hexakisphosphatecan prevent the disruption of the two mineralsand the formation of aluminium-phosphate salts,in contrast to the phosphate ion, which can dis-place silicon from the clay structures (Rajan,1975) and then progressively change the sitesavailable for adsorption, hampering the attain-ment of true equilibrium. The adsorptionmechanism, as hypothesized from the myo-inositolhexakisphosphate/phosphate ratio, suggests thatthree phosphate groups interact with the surfaceof both illite and kaolinite, while the other threephosphate groups remain free. However, it ispossible that the occupation of adsorption sites ispartly hindered by the organic moiety of theorganic phosphate. Possibly, if the phyllosilicatesare coagulated in an edge-to-face structure, myo-inositol hexakisphosphate adsorption is ham-pered and the number of phosphate groups thatremain free is greater than expected.

Finally, organic matter in both humic andnon-humic forms can participate in the retentionof inositol phosphates in soil (Hong and Yamane,1980, 1981; Borie et al., 1989; Makarov et al.,1997). This can occur through physical or chem-ical incorporation in the organic matter fraction,direct adsorption on the organic surfaces orindirect adsorption through polyvalent cationsthat act as bridges to form ternary organic mat-ter–metal– inositol phosphate complexes. Incor-

0.00

1.03106

2.03106

3.03106

4.03106

0.20

Goethite

KL

(I/m

ol)

Illite

Kaolinite

myo -Inositol hexakisphosphate/phosphate ratio

Fh−KGa2

Ferrihydrite

0.40 0.60 0.80

Fig. 13.4. Relationship between the Langmuir constants (KL) of goethite, kaolinite, illite, ferrihydrite andferrihydrite–kaolinite systems (Fh–KGa2) vs. the myo-inositol hexakisphosphate/phosphate molar ratio.(From Celi et al., 2003.)

Table 13.3. Values of the apparent associationconstants (1/mM) determined at 20°C in 50 mMpH buffer, containing myo-inositol hexakisphosphateat a concentration of 3.04 mM (pH 4.8, 6.0, 7.2)and 0.15 mM (pH 8.4, 9.4, 10.4). (From Graf,1983.)

pH K1 K2 K3

4.8 2.89 0.34 0.206.0 12.1 2.5 0.607.2 22.7 22.7 22.78.4 >1000 >1000 >10009.4 >1000 >1000 >1000

10.4 >1000 >1000 >1000

poration of inositol phosphates into the organicmatter structure by the formation of covalentbonds has also been hypothesized (Brannon andSommers, 1985), although this requires furtherinvestigation.

Complexation, Precipitationand Mineral Dissolution

In addition to adsorption onto solid surfaces, thestabilization of inositol phosphates in soil can berelated to their high capacity to complex metalcations (Cosgrove, 1980; Nolan and Duffin,1987). The order of stability of complexes withmyo-inositol hexakisphosphate is copper(II) > zinc> nickel(II) > cobalt(II) > manganese(II) >iron(III) > calcium (Martin and Evans, 1987;Nolan and Duffin, 1987). These complexes canhave many stability constants and become solu-ble at low concentrations or in certain pH ranges(Table 13.3), but as the concentration and pHincrease, they become less soluble and can pre-cipitate as insoluble salts (Martin and Evans,1987). Calcium phytate precipitation as a func-tion of pH and phytate activity occurs even atlow pH and can be responsible for a considerableproportion of the loss of inositol phosphates fromsolution (Celi et al., 2001a).

The ability of inositol phosphates to chelatecations can have important consequences for soilprocesses, because it affects the extent of sorp-tion, changes the speciation of inositol phos-phates and the relative composition of organicphosphorus in soil, enhances mineral weatheringand may have environmental implications. Forinstance, interaction of inositol phosphates withiron(III) was reported to transform labile organicphosphorus in manure applied to paddy soils intomore resistant forms, due to formation of insolu-ble iron-phytate (Zhang et al., 1994). The highstability of the cation–inositol phosphate com-plexes can cause the dissolution of minerals bydetaching metals from the surfaces. The forma-tion of insoluble salts can further enhance thisprocess by removing the metal ion from the reac-tion equilibrium. In fact, the formation of cal-cium complexes can cause dissolution of calcite(Celi et al., 2000), and the desorption of inositolphosphates from ferrihydrite–kaolinite mixed sys-tems and, to a lesser extent, from ferrihydrite and

goethite is followed by a consistent release of iron(Celi et al., 2003; Martin et al., 2004).

The ability of inositol phosphates to complexmetals has received great attention in the medicaland biological fields, because the anion can causemetal deficiency, especially in animals with dietsrich in seeds (Maga, 1982; Frossard et al., 2000,see Raboy, Chapter 8, this volume). Inositol phos-phates have been used in an immobilized form,bound to polyvinylpyridine, for removing heavymetal ions from the solution, thus offering apotential mechanism for decontaminating indus-trial or mining waste waters (Tsao et al., 1997).

Desorption of Inositol Phosphatesfrom Soil

Once sorbed in soil, inositol phosphates are notreadily released back to solution. Desorption ofmyo-inositol hexakisphosphate from iron oxides isa slow reaction that is affected by solution pH(Cabrera et al., 1981; Celi et al., 2003; Martinet al., 2004) and by the degree of phosphorus sat-uration (Parfitt, 1979; He et al., 1991, 1994;Martin et al., 2002). The configuration of thephosphorus–mineral complex and the formationof the multiple site-bindings play an importantrole in the strength of the bond and can furtherreduce the extent of desorption. Thus, no releaseof inositol phosphate was observed from goethite(Martin et al., 2004), it was negligible from ferri-hydrite, whereas it reached 16% of the adsorbedamount at basic pH from ferrihydrite–kaolinitemixed systems (Fig. 13.5; Celi et al., 2003).

Desorption of inositol phosphates boundto iron(III) oxides could increase under anaer-obic conditions following reduction to iron(II)(Schwertmann, 1991). This may explain the rapiddecomposition of inositol phosphates in anaerobicmarine sediments (Suzumura and Kamatani, 1995)and the absence of inositol phosphates in wetlandsoils subjected to anaerobic conditions for most ofthe year (Turner and Newman, 2005). However, itshould be considered that inositol phosphates couldre-precipitate with reactive amorphous iron oxyhy-droxides, as observed for inorganic phosphate (Sahet al., 1989). Moreover, it was suggested that thereduction of Fe(OOH)-phytate resulted in the for-mation of iron complexes and then insoluble Fe4-phytate (De Groot and Golterman, 1993).



Desorption of inositol phosphates from ironoxides is reported to be also affected by the pres-ence of competing ligands such as phosphate, cit-rate, oxalate and carbonate (Nagarajah et al.,1968; He et al., 1991; Presta et al., 2000; Martinet al., 2004). Other factors, such as the time fordesorption, solution/soil ratio and temperature,influence desorption of inorganic phosphate fromsoils and minerals (Barrow and Shaw, 1975;Barrow, 1983), but have not been systematicallyinvestigated for inositol phosphates.

Effects of Solution Characteristicson Abiotic Processes

The interaction of myo-inositol hexakisphosphatein soil involves adsorbents with variable chargesurfaces and an adsorbate with 12 ionizable –OHgroups. The pKa values are reported in Table13.4. The process is therefore affected by thecharacteristics of the soil solution, including pH,ionic strength, the nature and concentrationof electrolytes and the presence of competinganions.

Solution pH affects inositol phosphateadsorption by influencing both the charge of thereacting surfaces (Barrow et al., 1980; Bolan et al.,1986; Barrow, 1993) and that of the adsorbate,with a change in the relative concentrations ofthe anionic forms (Fig. 13.6). As the negativecharge on both adsorbate and adsorbent tends toincrease with pH, there is a reduction in theextent of myo-inositol hexakisphosphate sorptionat high pH in soils and on minerals with variablecharge surfaces (Anderson and Arlidge, 1962;Anderson et al., 1974; Shang et al., 1992; Celiet al., 2001a). Although HPO4

2− expresses agreater affinity than H2PO4

−, and similarly myo-inositol hexakisphosphate with eight chargesexpresses a greater affinity than the form with sixcharges, the increasing predominance of thesemore reactive forms with increasing pH appearsinsufficient to overcome the repulsive forcesraised by the increasing negative charge at theabsorbate surface.

At low pH the adsorption of myo-inositolhexakisphosphate on goethite is more pronouncedthan for phosphate (Fig. 13.7; Celi et al., 2001a),probably due to a different arrangement of themolecule on the oxide surface or the formation of

pH 3.5pH 4.5pH 7.0pH 8.5

Ferrihydrite Fh−KGa2 Goethite

myo-IP6myo-IP6myo-IP6 PiPi Pi0

5

10

Des

orbe

d P

(%

)

15

20

25

30

Fig. 13.5. Desorption of myo-inositol hexakisphosphate (myo-IP6) and phosphate (Pi) from ferrihydrite,ferrihydrite–kaolinite systems (Fh–KGa2) and goethite, as affected by pH. (Adapted from Celi et al.,2003; Martin et al., 2004.)


insoluble iron-phytate salts if the pH is lowenough to cause mineral dissolution (Andersonand Arlidge, 1962; Anderson et al., 1974). Withincreasing pH the behaviour is opposite, with amore pronounced decrease in adsorption capacityfor the organic compound (Anderson et al., 1974;Shang et al., 1992; Celi et al., 2001a). This isattributed to the reduced capacity of myo-inositolhexakisphosphate compared to phosphate to neu-tralize the hydroxyl ions released from the surfaceduring adsorption (Table 13.4; Shang et al., 1990;Celi et al., 2001a).

The decrease in adsorption with increasingpH can facilitate mineralization of inositol phos-phates in soil at near-neutral pH (Dalal, 1977),which could account for the greater accumula-tion of inositol phosphates in acid rather than

alkaline soils (Turner et al., 2002; see Turner,Chapter 12, this volume). This is also related tothe optimal conditions for microbial activity,although phytase activity is optimum nearer topH 5 (Ullah and Gibson, 1987). The sorption ofphytase in soils is discussed elsewhere in this vol-ume (see George et al., Chapter 14).

In addition to pH, the electrical charge ofthe adsorbent is affected by the nature and con-centration of electrolytes concentrating in thedouble layer surrounding the charged particles(van Olphen, 1977; Barrow et al., 1980; Bowdenet al., 1980; Barrow, 1993). With monovalentcations the adsorption is only slightly affected bythe concentration of electrolytes. As this concen-tration increases, the adsorption of inositol phos-phates should decrease at pH values lower than

Table 13.4. Dissociation acid constants (pK) of myo-inositol hexakisphosphate (myo-IP6) (from Costelloet al., 1976) and H3PO4 (from Corbridge, 1985).

Molecule pK1 pK2,3 pK4–6 pK7 pK8 pK9 pK10,11 pK12

myo-IP6a 1.1 1.5 1.8 5.7 6.9 7.6 10.0 12.0

H3PO4 2.0 6.8 12.3 − − − − −

amyo-Inositol hexakisphosphate = C6H6(H2PO4)6.

20

20

40

60

Con

cent

ratio

n (%

) 80

100

120

3 4 5 6

pH

7 8 9 10

HPO42−

IHP8−

IHP6−

H2PO4−

Fig. 13.6. Aqueous speciation of myo-inositol hexakisphosphate (IHP) and inorganic phosphate atincreasing pH. (From Celi et al., 2001a.)


the point of zero charge of the mineral andincrease at pH values higher than the point ofzero charge. This would be caused by a reduc-tion of the absolute value of the electric potentialat the shear plane, due to compression of thedouble layer, although no studies have directlyaddressed this.

Surface charge is strongly affected by thepresence of polyvalent cations. For instance in thepresence of calcium, the surface of goethiteremained positive even at high pH (9–10) and inthe presence of low concentrations of electrolyte(Celi et al., 1998). The ability of inositol phos-phates to form complexes with polyelectrolytescould further favour the interaction with adsor-bates by the formation of bridges or salts that pre-cipitate on reacting surfaces. With calcium, theadsorption of myo-inositol hexakisphosphateincreases above pH 5, well beyond its capacity toform a monolayer on goethite, due to the simulta-neous occurrence of adsorption and precipitationof insoluble calcium-phytates (Celi et al., 2001a).

The adsorption of inositol phosphates onsoil components is so strong that the competitionof other ligands in the bulk solution for the samesites of adsorption is expressed slightly, asobserved with phosphate, citrate or carbonate(Anderson et al., 1974; Presta et al., 2000; Martinet al., 2004). Conversely, myo-inositol hexakisphos-

phate can displace phosphate from mineralsurfaces, either before or during treatment withthe latter, and inhibit phosphate adsorption(Anderson et al., 1974; De Groot and Golterman,1993; Presta et al., 2000). The release of phos-phate and organic matter into solution upon myo-inositol hexakisphosphate addition, and theinhibition of their re-sorption, have also beenobserved in soils (Anderson et al., 1974; De Grootand Golterman, 1993; Leytem et al., 2002) andcould accelerate phosphorus transfer to waterbodies in runoff (see also Leytem and Maguire,Chapter 10, this volume).

Effects of Inositol PhosphateSorption on Surface Properties

The adsorption of high charge-density anions oncolloidal particles creates new surfaces with a dif-ferent charge and electric potential, thus affectingtheir dispersion/flocculation behaviour. Theadsorption of myo-inositol hexakisphosphate ondifferent iron oxides and phyllosilicates canreverse the initial net positive charge of the sur-faces, thus increasing particle–particle repulsiveforces and colloidal dispersion (Celi et al., 1999,2003). This is attributable to the phosphate

1086pH

420.0

1.0

2.0

Qa

(µm

ol P

/m2 )

3.0

4.0

5.0

Fig. 13.7. Effect of the pH on sorption (Qa) of myo-inositol hexakisphosphate (shaded symbols) andphosphate (open symbols) by goethite in 0.01 M KCl. (From Celi et al., 2001a.)

groups of myo-inositol hexakisphosphate that arenot involved in the binding mechanism and thathave hydroxyl groups that dissociate at pH > 2(Table 13.4). The new surface will have a highercharge when fewer phosphate groups are bound;thus, phyllosilicate complexes with myo-inositolhexakisphosphate present a larger charge thaniron oxide complexes (Celi et al., 1999). The over-all net negative charge of the surface is reachedwith only low concentrations of inositol phos-phate and over a large range of pH, whereas withinorganic phosphate the negative charge isobtained only at pH > 5 and with a high per-centage of phosphorus coverage.

Monovalent cations in the bulk soil solutionaffect the changes in surface charge only in termsof absolute values, whereas with polyvalent cationsthe surface charge remains positive due to the for-mation of mineral–inositol phosphate–cation com-plexes that counterbalance the effect of the organicanion (Celi et al., 2001a).

The ability of inositol phosphates to detachcations from minerals, as shown for iron releasedfrom ferrihydrite (Celi et al., 2003), could haveimportant effects on the weathering of surfaceminerals, although few studies have been devotedto this topic. Moreover, in contrast to phosphate,the relatively large size of the inositol phosphatemolecule should preclude its diffusion into themineral pores through time, allowing a true equi-librium to be reached more rapidly than withinorganic phosphate.

Summary and Recommendationsfor Future Research

Abiotic reactions are the main processes stabiliz-ing inositol phosphates in soil and limiting theirdegradation by plants and microorganisms. Theaffinity of these phosphate monoesters for claysand metal oxide surfaces, their ability to formcomplexes with polyvalent cations and insolublesalts, and their incorporation in organic struc-tures account for the accumulation of inositolphosphates compared to other organic phospho-rus compounds in soil. Recent studies haveadvanced our understanding of the interaction ofinositol phosphates with pure or more complexminerals and organic matter, although someaspects remain unknown. In particular, future

studies should address the effects of temperature,solution/soil ratio, concentration of electrolytesand stereoisomeric forms of inositol phosphatesother than myo-, on the extent and mechanism ofinositol phosphate adsorption. Moreover, therole of organic matter should be expanded, andattention paid to the influence of reaction kineticson the long-term fate of sorbed inositol phos-phates. A more comprehensive investigationof the processes regulating inositol phosphatebehaviour under anaerobic conditions is alsonecessary to understand the potential bioavail-ability of inositol phosphates under changingredox conditions.

The dispersal of colloidal particles followinginositol phosphate adsorption on minerals hasimportant environmental implications (seeLeytem and Maguire, Chapter 10, this volume).Although adsorption immobilizes a large amountof inositol phosphate in soil, there is a greatpotential for transfer to water bodies in runoff asparticulate phosphorus (Turner, 2005). The dis-persion caused by adsorption of inositol phos-phates can dramatically affect the transport ofcolloids in soil and may explain the presence ofinositol phosphates in the particulate form foundin rivers and lakes (McKelvie et al., 1995;Suzumura and Kamatani, 1995). This can con-tribute to eutrophication, which is currently amajor threat to global water quality (Correll,1998; Turner et al., 2002; see McKelvie, Chapter16, this volume).

Similarly, the ability of inositol phosphatesto detach metals from minerals, together with thepotential dispersion of particles upon inositolphosphate adsorption, could enhance mineralweathering and clay or metal translocationthrough the soil profile, with important effects onpedogenic evolution.

In the future, integration of research on abiotic and biotic processes should improve ourability to evaluate the availability of inositol phos-phates to plants and their transport to water bod-ies. Studies should also include other soil organicphosphorus compounds. Integrating this informa-tion with the large body of literature devoted toinorganic phosphate will enable a comprehensiveunderstanding of the terrestrial phosphorus cycle,and contribute to the development of land man-agement strategies that combine agronomic pro-ductivity with sustainable management of theenvironment.



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phosphate by variable charge surface. Australian Journal of Soil Research 18, 395–404.Bolan, N.S., Syers, J.K. and Tillman, R.W. (1986) Ionic strength effects on surface charge and adsorption of phos-

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14 Interactions Between Phytases andSoil Constituents: Implications for the

Hydrolysis of Inositol Phosphates

Timothy S. George1, Hervé Quiquampoix2, Richard J. Simpson3

and Alan E. Richardson3

1Scottish Crops Research Institute, Invergowrie, Dundee DD2 5DA, UK; 2Unité deScience du Sol, INRA-ENSAM, 2 Place Pierre Viala, 34060 Montpellier Cedex 1,

France; 3CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia

A large proportion (up to 80%) of soil phosphorusoccurs in organic forms (Harrison, 1987), ofwhich derivatives of inositol phosphates constitutea considerable fraction (Anderson, 1980; seeTurner, Chapter 12, this volume). The bioavail-ability of inositol phosphates depends on theirmineralization by phytases (myo-inositol-hexak-isphosphate phosphatases), which come in severalclasses (EC 3.1.3.8, EC 3.1.3.26 and EC 3.1.3.72),that initially cleave phosphate at different posi-tions on the myo-inositol ring (see Mullaney andUllah, Chapter 7, this volume). Phytases were firstrecognized almost a century ago (Suzuki et al.,1907; Dox and Golden, 1911) and have manybiological sources, including plants, animals and awide range of microorganisms (Table 14.1). Theyare particularly important in the soil environ-ment, having been identified in plant roots, fungi,yeasts and bacteria (Irving, 1980).

Phytases do not constitute a major componentof plant root exudates and appear to be absentfrom monogastric animal digestive systems (Hayeset al., 1999; Brinch-Pedersen et al., 2002).Therefore, research into the role of phytases in bio-logical phosphorus cycling in both natural andagricultural systems has focused primarily on phy-tases produced by microorganisms. Both soil bacte-ria and fungi produce extracellular phytases, which

give plants a nutritional benefit when present in therhizosphere (Findenegg and Nelemans, 1993;Richardson et al., 2001b; Idriss et al., 2002).Microbial phytases have also been specifically engi-neered for supplementation of monogastric animalfeeds (Lehmann et al., 2000) and have the potentialto enter soil through animal excreta. In recentyears, transgenic plants that express microbial phy-tase genes have been produced for use in animaldiets (Pen et al., 1993) and have been evaluated fortheir capacity to improve plant nutrition(Richardson et al., 2001a; see Richardson et al.,Chapter 15, this volume). Transgenic animals withenhanced phytase activity in saliva have also beendeveloped (Golovan et al., 2001). Of particular sig-nificance to the mineralization of inositol phos-phates in the soil environment is the expression offungal (Richardson et al., 2001a; Zimmermannet al., 2003) and bacterial (Lung et al., 2005) phytasegenes in plants. These plants show improved phos-phorus nutrition when grown under controlledconditions, but this is compromised when plantsare grown in the more complex soil environment(George et al., 2004, 2005a,c).

Whilst mobility of both substrate (inositolphosphates) and product (phosphate) of the phy-tase reaction are likely to be major limitations tothe efficacy of phytases in the soil environment

222 T.S. George et al.

(see Celi and Barberis, Chapter 13, this volume),edaphic factors directly affecting the stability andcatalytic efficiency of phytases are also important(George et al., 2005b, 2006). In this chapter wereview the interactions of phytases with soil con-stituents and consider the implications of this forthe hydrolysis of inositol phosphates in soil.Although many edaphic factors alter the synthesisand secretion of phytases through direct impactson biological systems, this review focuses onimpacts of the soil environment on the stabilityand catalytic efficiency of discrete phytase pro-teins following their release from the cytoplasm.

Factors Affecting Phytase Activity in Soil

Most of the phytases produced by soil organismsare thought to be released as extracellularenzymes by active exudation (Tarafdar et al., 2002)and thus have only a short contact time with thecytoplasmic environment. However, some of the

activity found in soil will presumably be passivelyreleased following cell lysis and thus be adaptedspecifically to the intracellular environment. Oncein soil, phytases must withstand many factors inorder to remain functional (Fig. 14.1). Theseinclude (Goldstein, 1976; Gianfreda and Bollag,1996; Nannipieri et al., 1996; Nannipieri andGianfreda, 1998):

1. deactivation and inhibition by adsorption andimmobilization on soil solid particles;2. proteolytic and microbial mediated degrada-tion;3. inhibition by interaction with metal ions,anions and metabolites; and4. denaturation by soil environmental factors(temperature, pH, water content, light).

The impact of these factors on phytases found inthe soil environment is potentially large (Fig. 14.1),leaving little phytase activity for longer-termhydrolysis of inositol phosphates. Few studies haveexplicitly measured phytase activity in soil, but‘baseline’ activities, if detectable, are in the range of10–300 pKat/g soil ( Jackman and Black, 1952;

Table 14.1. Biochemical properties of a selection of phytase enzymes from a range of biologicalsources. (Adapted from Vats and Banerjee, 2004.)

Molecular Isoelectric pH Temperature Source of phytase weight (kDa) point (pl) optimum optimum (°C) Reference

Bacteria

Bacillus sp. DS 11 44 5.3 7.0 70 Kim et al. (1998a,b)Bacillus subtilis 36–38 6.3 6.0–6.5 60 Kim et al. (1999)B. licheniformis 44, 47 5.0, 5.1 4.5–6.0 55–65 Kerovuo et al. (1998,

2000)Escherichia coli 42 6.3–6.5 4.5 60 Greiner et al. (1993)

Yeast

Saccharomyces 120 – 2.0–2.5 55–60 Han et al. (1999)cerevisiae

Fungi

Aspergillus niger 85 4.5 2.5, 5.0 58 Ullah and Gibson (1987)(phyA)

A. niger (phyB) 68 4.0 2.5 63 Erlich et al. (1993); Ullah and Cummins (1987)

A. oryzae 120 4.2 5.5 50 Shimuzu (1993)A. fumigatus 91 7.3 55 Wyss et al. (1999b)Peniophora lycii 72 3.6 4.0–4.5 50–55 Lassen et al. (2001)

(phyA)

Plants

Glycine max 60 5.5 4.5–4.8 55 Gibson and Ullah (1988)

Interactions Between Phytase and Soil Constituents 223

T.S. George et al., 2002 unpublished data), with themaximum represented by a mor humus layer froma spruce woodland (Svenson, 1986). In comparisonwith the activity of enzymes released to the soilthrough various biological processes ( Jackman andBlack, 1952), these activities are small. For exam-ple, samples taken from the rhizosphere of trans-genic plants, which exude large amounts ofAspergillus niger phytase, had activities against inosi-tol phosphate representing less than 1% of thatknown to be exuded to the soil by the plants(George et al., 2004, 2005b). Moreover, baselinephytase activity in soil appears insignificant whencompared with total phosphomonoesterase activi-ties, which are 1–2 orders of magnitude greater,but is similar to those of soil phosphodiesterases(Eivazi and Tabatabai, 1977). In spite of this, toler-ance of phosphatases (including phytases) to theextracellular environment will vary depending onbiochemical properties of the enzyme (Table 14.1),which will have an important influence on thebaseline activity in soil and thus to the biologicalcycling of inositol phosphates.

Interaction of phytases with the soil solid phase

Proteins have an affinity for the interfacebetween the aqueous and solid phase of soil, so

adsorption of enzymes is common (Norde andLyklema, 1991). In some cases this can inhibitenzyme activity irreversibly (Quiquampoix,1987a, 2000; Quiquampoix and Mousain, 2005).Adsorption of phytases may reduce the affinityfor substrates and thus reduce the effective activ-ity. However, immobilization protects phytasesfrom degradation (Naidja et al., 2000) and may beresponsible for their long-term persistence in soil(Nannipieri et al., 1996).

Processes of immobilization and adsorption

Phytase from A. niger was rapidly sorbed whenadded to a range of soils with varying adsorptioncapacity (George et al., 2005b). However, most ofthe activity was immediately recovered on thesoil solid phase. This indicates that the initial fateof this particular phytase upon introduction tosoil was immobilization by adsorption to soil solidconstituents. This may involve binding to solidsupports by covalent bonds, cooperative adsorp-tive interactions, and entrapment and encapsula-tion in stable aggregates (Gianfreda and Bollag,1996). Electrostatic and van der Waals forces, aswell as hydrophobic interactions, have been sug-gested for adsorption to clays (Quiquampoix,2000; Quiquampoix et al., 2002), and intercala-tion of proteins in layered clays may also occur.In contrast, ion exchange, entrapment in organicnetworks and covalent bonds may account

Plant

Bacteria

Fungus

Yeast

Manuresanddiet

supplementSou

rces

of p

hyta

se in

put t

o so

il

Loss ofPhytase activity with time

Proteolysis

MicrobialdegradationAdsorption

Interaction with ions

Soil environment− pH− temperature− water

− light

Soilsolid

Soilsolution

Endogenous

Plant residues

Inositolphosphates

Microbial

Manures

Soilphytase

Fig. 14.1. Sources and fate of phytases in the soil environment. The schematic demonstrates that thebiological origin of phytases in soil is varied and that phytase activity may be lost through a wide rangeof competing biological, chemical and physical processes before its interaction with inositol phosphates,either within the soil solution or at the boundary with the solid phase.

for the stable association between phytases andhumic materials (Perez-Mateos et al., 1991;Gianfreda and Bollag, 1996). In addition tocharged amino-acid residues, protein groupsthat are potentially free to react with soil con-stituents include carboxyl, phenol, thiol, aliphatichydroxyl and amide groups (Brown andHasselberger, 1971). The heterogeneous natureof proteins also means they contain both polarand non-polar domains that can interact withhydrophylic and hydrophobic surfaces in the soil,respectively (Laidler and Sundaram, 1971).

Soils are also extremely heterogeneous.They contain a mixture of organic materials ofvarious levels of decomposition, clay and metal(hydr)oxide minerals, and combinations of thesematerials, which produce complexes with a widerange of adsorption properties. Furthermore,enzymes can be adsorbed to live biological mate-rial such as cell walls and mucigel (Ciurli et al.,1996; Marzadori et al., 1998). Extracellularenzymes in soil are primarily associated withhigh-molecular weight organic matter and clay-sized particles, including specific clay mineralsand humified organic matter (Kanazawa andFilip, 1986; Perez-Mateos and Rad, 1989; Rojoet al., 1990; Marx et al., 2005). However, detailedcytochemical and microscopy techniques havebeen unable to verify the presence of discreteenzyme–clay complexes (reviewed by Ladd et al.,1996). While coarse organic matter and fine clayfractions are likely to be the main sites foradsorption and immobilization of phytases, theyare also the primary habitats for microorganisms,the major source of phytases in soil (Rojo et al.,1990; Marx et al., 2005).

Despite the heterogeneity of soil, predictablepatterns of biological and biochemical activityare evident. For example, biological activity andits products decline with depth, which has impli-cations for the distribution of phytase activity interms of its production and the availability oforganic sites for immobilization (Nannipieri andGianfreda, 1998). Moreover, the major zone ofconcentration of phytases in soil is likely to be therhizosphere, where plant and soil are in contactand microorganisms are abundant (Tinker andNye, 2000). As such, the rhizosphere will likely bethe initial external environment experienced bymany extracellular phytases.

Clay minerals will provide a major propor-tion of the surface area for adsorption of phytases

in many soils. The adsorption capacity of the clayfraction depends on the type of clay present (e.g.layered clays vs. non-layered clays, permanentcharge vs. variable charge minerals) with majordifferences being exemplified by comparisonsbetween montmorillonite and kaolinite, the for-mer being the stronger adsorbate. While much ofthe theoretical research on adsorption of enzymesby clays has involved pure forms of montmoril-lonite (Quiquampoix and Mousain, 2005), suchmodel systems may be poorly representative ofthe soil environment. ‘Perfect clays’ rarely exist insoil, where magnesium, aluminium and silicon inthe mineral lattice are commonly substituted forby impurities, causing changes in the electricalcharge of the clay (Rowell, 1994).

Dispersion of clays by large concentrationsof certain cations such as sodium and potassiumcan also modify sorption surfaces and affectenzyme adsorption (Violante and Gianfreda,2000). In addition, clays are often coated withorganic material or metal (hydr)oxides, whichalters their adsorption capacity. For example,enzymes are less adsorbed on aluminium-coatedmontmorillonite than on pure clay (Violante andGianfreda, 2000). Even the presence of phytases(and other proteins) may alter the adsorptionenvironment presented by the clay, as rapidunfolding of the enzyme at the clay surface mayproduce a denatured protein monolayer coveringthe clay particle, acting as a more benign adsorp-tion environment for subsequent enzymes (Brownand Hasselberger, 1971).

In addition to interaction with mineral mate-rials, phytases also form complexes with humicsubstances and their constituents, including phenolsand quinones (Ladd and Butler, 1975; Wetzel,1993; Gianfreda and Bollag, 1996). Reactions canbe reversible (e.g. hydrogen bonding between phe-nolic groups and oxygen in the peptide bond) orirreversible (e.g. covalent bonding between termi-nal amino and sulphhydryl groups of the enzyme)(Ladd and Butler, 1975). Further interaction withother soil components may also occur; for example,flocculation of organic–protein complexes wasenhanced by the presence of polyvalent cations andclay minerals (Rao and Gianfreda, 2000; Violanteand Gianfreda, 2000).

Phytases added to soil collected from therhizosphere of plants were less rapidly adsorbedto the solid phase than when added to soilthat had not been affected by the presence of a



growing plant root (George et al., 2005b). In asoil with relatively little sorption capacity, phy-tases from A. niger remained active in thesolution phase to a greater extent in rhizospheresoil than in bulk soil. In contrast, in soil withgreater clay and organic matter content andlarger cation–anion exchange capacity, a smalleramount of phytases remained in solution underrhizosphere conditions. Such differences arelikely to be associated with differences in thechemistry and biochemistry of rhizosphere andbulk soil, such as pH and the presence of proteinsand organic anions. The presence of organicanions, such as citrate and malate, has beenshown to increase the amount of acid phos-phatase retained in soil solution (Huang et al.,2002, 2003) due to ligand exchange reactions ofthese anions in the presence of metal (hydr)oxidesurfaces (Violante and Gianfreda, 2000). Giventhe heterogeneous charge characteristics of pro-teins (including phytases), competitive exchangereactions will involve anions as well, while otherhetero-valent proteins may be of similar impor-tance. The significance of electrostatic interac-tions between phytases and soil constituents willalso depend on the ionic strength of the soilsolution, as electrostatic forces diminish withincreasing salt concentration (Goldstein, 1976).Differences in ionic strength may be especiallyimportant in determining retention of phytaseswhere soil moisture fluctuates and salinity andsodicity are apparent.

Soil pH exerts a strong control on phytaseadsorption. Adsorption of an A. niger phytase to arange of soils was shown to be complete at pH4.5, close to the isoelectric point of this enzyme(pH 4.8–5.2; Wyss et al., 1999a). In contrast, theproportion of phytase activity recovered in soilsolution was greater as pH was increased(Fig. 14.2). A number of enzymes have maxi-mum adsorption on a range of different surfacesat their isoelectric point (Kondo et al., 1993;Quiquampoix et al., 1993, 2002; Violante et al.,1995; Huang et al., 1999). Proteins at their iso-electric point have no net electric charge, andtherefore only weak interactions such as van derWaals or hydrogen bonding are invoked for suchadsorption (Violante and Gianfreda, 2000).Nevertheless, entropic factors, such as hydropho-bic interactions or modification of proteinconformation towards a less-ordered secondarystructure, can result in stronger interactions with

soil constituents, even at a protein’s isoelectricpoint. The general partitioning of phytase activ-ity to the solution phase with increased pH isattributable to greater electrostatic repulsionabove the isoelectric point of the protein dueto enthalpic forces (Kondo and Higashitani,1992; Kondo et al., 1993; Quiquampoix et al.,1993).

To investigate further the importance ofthe isoelectric point for adsorption of phytases,the adsorption of Peniophora lycii phytase (isoelec-tric point ≈ 3.6) and that produced by A. niger(isoelectric point ≈ 4.8) were compared follow-ing addition to a range of soils (George et al.,2006). It was shown that P. lycii phytaseremained active in solution at a soil pH of ~5.5,whereas A. niger phytase was rapidly adsorbedto soil solid phase (Fig. 14.3). This is poten-tially important as the known range of isoelec-tric points (pH 3.6–7.3) for phytases is large(Table 14.1), suggesting that different enzymeswill be more or less adsorbed over a wide rangeof soil pH.

Biochemical differences in proteins havealso been invoked in studies of the adsorption ofphosphatases from ectomycorrhizal fungi, withinterspecific and intraspecific differences inadsorption properties being observed for differentenzymes (Quiquampoix and Mousain, 2005).Phosphatases from Pisolithus tinctorius showed noadsorption on montmorillonite between pH 2and 8, whereas phosphatases from other species(Cenococcum geophilum, Hebeloma cylindrosporum)showed an increasing adsorption from pH 6 to 4,followed by complete inhibition of the catalyticactivity of the adsorbed fraction. Furthermore,adsorption of phosphatase from Suillus bellini var-ied with pH, whereas that from S. mediterraneensiswas completely adsorbed across a range of pH,although catalytic activity of both enzymes wasmaintained regardless of their adsorption(Quiquampoix and Mousain, 2005). It is there-fore apparent that not only does specific variabil-ity in the adsorption of distinct phytases occur,but differences in inhibition by adsorption arealso probable. Differences in biochemical charac-teristics of phytases will affect their mobilityand are therefore potentially important for theinteraction of phytases with inositol phosphates(Wyss et al., 1999a; Vats and Banerjee, 2004;Quiquampoix and Mousain, 2005; George et al.,2006).


Inhibition and protection of phytasesby immobilization

Protection of enzymes has been demonstratedupon adsorption to a number of soil surfacesincluding clay, metal (hydr) oxides, organic mate-rial and mucigel produced by plants and microor-ganisms (Rao et al., 1994; Ciurli et al., 1996; Naidjaet al., 2000). Immobilized enzymes usually showincreased resistance to temperature, proteaseactivity and microbial degradation (Estermannet al., 1959; Makboul and Ottow, 1979; Sarkaret al., 1989; Kandeler, 1990; Perez-Mateos et al.,

1991; Nannipieri, et al., 1996; Naidja et al., 2000;Violante and Gianfreda, 2000) and improvedresistance to freezing and thawing, wetting anddrying, changes in pH and presence of heavy met-als (Gianfreda and Bollag, 1996).

Once adsorbed, A. niger phytases added to arange of soils showed greater stability thanenzymes not in the presence of soil (George et al.,2005b, 2006). Moreover, the activity of endoge-nous soil phosphatases appears to be more stablethan newly added or recently immobilizedenzymes, presumably due to persistence of

pH

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L.S.D.(P < 0.05)

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(i) (ii)

(a)

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Fig. 14.2. The effect of pH on the adsorption of Aspergillus niger phytase in soil. Phytase activity,measured as a proportion (%) of the amount initially added to soil, is shown (i) in the solution phase and(ii) on the solid phase of two soil types: (a) a Spodosol and (b) an Alfisol. Phytase activity against myo-inositol hexakisphosphate was measured at three time points (1 h, 48 h and 8 days) after addition ofphytase to soil suspensions buffered at a range of pH 4.5–8.5. Data show the mean of three replicateswith bars representing two standard errors. Differences between enzyme activity measured at differentpH and at different times (within each soil type) for both solution and solid phase were established usingANOVA. Least significant difference (P < 0.05) is presented as a bar for each soil type and phase withinsoil. (From George et al., 2005b.)

stabilized forms of the enzymes over time (Perez-Mateos et al., 1991). The resilience of immobilizedphytases may be due to a number of factorsincluding: (i) concurrent adsorption of proteasesand inhibitory substances (metal ions, chelators,etc.) (Ciurli et al., 1996; Demanèche et al., 2001;Casucci et al., 2003; Renella et al., 2003); (ii) sterichindrance against relatively large proteases andmicroorganisms for enzymes embedded inorganic matrices (Wetzel, 1993); and (iii) confor-

mational changes that increase the stability ofthe protein structure, preventing autolysis andincreasing the energy required for denaturation(Quiquampoix and Mousain, 2005).

Despite the possibility of protection of a pro-portion of phytase activity by adsorption, muchactivity may also be inhibited by interaction withsoil solid surfaces (George et al., 2005b).Generally, adsorption in mixed environmentssuch as soil is less inhibitory to enzyme activitythan adsorption to pure clays or some organicmaterials. For example, several studies demon-strated less inhibition of phosphatase activitywhen adsorbed to montmorillonite coated withaluminium hydroxides than on clean clay sur-faces (Rao et al., 1994; Geiger et al. 1998a; Bayanand Eivazi, 1999; Huang et al., 1999). Reducedinhibition has similarly been observed for alu-minium hydroxide–tannic acid complexes com-pared with tannic acid alone (Gianfreda et al.,1993).

Soil phosphatase activity is commonly corre-lated with organic matter content, whereas nega-tive relationships with clay content are oftenobserved (Harrison, 1983; Feller et al., 1994).Faster inhibition of phytase and other phos-phatases following adsorption has similarly beenshown to occur in soil with increasing dominanceof clay compared to organic matter (Sarkar et al.,1989; George et al., 2005b). The temporal pat-tern of phytase degradation following adsorptionto soil solid constituents (George et al., 2005b) wassimilar to those observed for acid phosphataseadsorbed to various clay–sesquioxide andorganic surfaces (Rao et al., 2000) and otherenzymes on mixed clay–sesquioxide–organicsurfaces (Nannipieri et al., 1996; Naidja et al.,2000). Interestingly, the temporal unfolding ofmodel proteins adsorbed on montmorillonite(Quiquampoix et al., 2002) also follows a similarpattern, suggesting a potential mechanism forphytase deactivation after adsorption to soil(Leprince and Quiquampoix, 1996). This is cor-roborated by the fact that loss of A. niger phytaseactivity on the soil solid phase is irreversible(George et al., 2005b). The rate of irreversibleinhibition after adsorption is also pH-dependent(Fig. 14.1) and a function of electrostatic forces.At pH below the isoelectric point of the phytase,increased conformational change and denatura-tion following adsorption would be expected(Leprince and Quiquampoix, 1996).


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Fig. 14.3. Adsorption of phytase from Aspergillusniger and Peniophora lycii as a function ofdiffering isoelectric points (pI). Activity (nKat/g soil)against myo-inositol hexakisphosphate of the twofungal phytases with different pI (4.8 and 3.6,respectively) was measured in the solution phaseof two soil types: (a) a Spodosol and (b) an Alfisol.Phytase activity was measured at time pointsbetween 1 and 24 h after the addition of phytasesto soil suspensions buffered at pH 5.5 and 7.5.Data show the mean of three replicates with barsrepresenting two standard errors. (From Georgeet al., 2006.)

The resilience of phytase in the soil environ-ment also depends on the type of clay mineralpresent; for example, kaolinite is less inhibitory toadsorbed enzymes than clay with interlaminarspaces, such as montmorillonite (Nannipieri et al.,1996). Moreover, the capacity of pure montmo-rillonite for adsorption and inhibition of phos-phatase is greater than intercalated equivalents,suggesting that phosphatases are absorbed andstrongly inhibited in the interlayer region of thepure clay (Kelleher et al., 2004).

The potential importance of organic matterin protecting phytase activity is implied by thefact that remnant phosphatase activities in soilare associated almost exclusively with organic,rather than clay, surfaces (Ladd et al., 1993). Theprotective effect of organic material has beenattributed to it being structurally diverse, withmacropores where enzymes can lodge and besterically protected from microorgansims andproteolytic enzymes, while still permitting accessby relatively low-molecular weight substrates(Estermann et al., 1959; Burns et al., 1972; Naidjaet al., 2000). In fact, the larger the humic–enzymecomplex is, the greater is the apparent resistanceof the immobilized enzyme to degradative factors(Nannipieri et al., 1988).

The organic portion of clay–humus com-plexes is considered to be important in protectingenzymes from the inhibitory effects of clay, byacting as a barrier between clay and enzyme(Quiquampoix, 1987b; Quiquampoix et al., 1995,2002). However, some organic compounds canalso be inhibitory to phosphatase activity. Forexample, activity is reduced when phosphatase iscomplexed with tannic acid (Rao and Gianfreda,2000) and phenolics (Wetzel, 1993), with inhibi-tion being caused by blocking of the enzymeactive site (Ladd and Butler, 1975). Para-doxically, phosphatase–tannic acid complexesformed in the presence of both iron-oxides andmontmorillonite retain greater activity than inthe presence of the tannic acid or minerals alone(Rao and Gianfreda, 2000).

Of particular interest is that the resilience ofphytase towards the adsorption environmentappears to be determined by whether theenzymes usually have extracellular or intracellu-lar function. For example, intracellular phytasesfrom wheat and S. collinitus were completelyadsorbed on clay particles across a range of pH,and showed complete inhibition of catalytic activ-

ity (Matumoto-Pintro and Quiquampoix, 1997).In comparison, two extracellular phytases fromA. niger and H. cylindrosporum retained significantcatalytic activity in the presence of clay, irrespec-tive of their degree of adsorption (Matumoto-Pintro and Quiquampoix, 1997). Similarly,extracellular phosphatase from maize roots wasless inhibited in soil than that from wheat germ(Dick et al., 1983). Such retention of activity byextracellular enzymes upon adsorption has beensuggested to be an evolutionary consequence ofan organism’s requirement for a functional extra-cellular enzyme (Quiquampoix, 2000). Indeed, ithas been suggested that adsorbed–uninhibitedenzymes may act as an indicator of transient con-centrations of biochemically available substratesto microorganisms in the surrounding soil niche(Burns, 1982; Allison and Vitousek, 2005).

Effects of phytase immobilization on reaction kinetics

The heterogeneity of the soil environment meansthat most extracellular phytase will not catalysereactions as efficiently as in homogeneous in vitrosystems. The kinetics of immobilized phytase insoil are likely to be different to that of free phy-tase in soil solution (Nannipieri and Gianfreda,1998). Studies with model phosphatase–clay andphosphatase–metal (hydr)oxide complexes show ageneral loss in enzyme activity due to declines inthe velocity of the reactions (Vmax) or a reductionin the affinity of the enzyme for its substrate(increased Km) (Gianfreda and Bollag, 1996;Huang et al., 1999; Quiquampoix and Mousain,2005). Similar declines in velocity and affinityhave also been noted with adsorption to modelmucigel compounds (calcium–polygalacturonate)(Marzadori et al., 1998) and whole soil (Perez-Mateos et al., 1991; Gianfreda and Bollag, 1994;George et al., 2006). However, mixtures of soilcomponents appear to have mitigating effects.For example, phosphatase associated with tannicacid had a reduced reaction velocity and sub-strate affinity, but when these complexes were inthe presence of montmorillonite, velocity wasunaffected and the affinity of the reaction wasincreased (Rao and Gianfreda, 2000).

Different clay types also have differentialeffects on the kinetics of reactions. Montmorillonitereduced the velocity of phosphatase reactions,while kaolinite reduced substrate affinity (Dick and


Tabatabai, 1987). Adsorption of phytase from A.ficuum (now known as A. niger) on gelatin particleshad similar effects of declining velocity and affinity(Liu et al., 1999). Moreover, when phytases fromtwo soil fungi (A. niger and P. lycii) were added tosoil, both their velocity and substrate affinitydeclined (George et al., 2006). Of particular interestwas that although velocity was not differentbetween the two phytases in the soil environment,the affinity for inositol phosphate of the phytase insolution (P. lycii) was double that of the adsorbed(A. niger) enzyme (George et al., 2006). This suggeststhat greater proportions of phytase in solutionincrease the capacity for interaction with inositolphosphates. This was further demonstrated by thegreater ability of P. lycii phytase, which is lessadsorbed to soil surfaces than A. niger phytase, tomineralize inositol phosphates endogenous to arange of soils (George et al., 2006). Interestingly, ithas also been demonstrated that the extracellularphytase from a range of soil fungi (Aspergillus spp.,Emericella spp. and Penicillium spp.) is more effec-tive at hydrolysing inositol phosphates comparedwith intracellular equivalents from the same organ-isms (Tarafdar et al., 2002), suggesting the existenceof traits peculiar to the extracellular protein thatallow more effective function in the soil environ-ment.

The observed effects of immobilization ofphytases on their catalytic activity may dependon the following factors (Laidler and Sundaram,1971; Ladd and Butler, 1975; Engasser andHorvath, 1976; Goldstein, 1976):

1. conformational change upon adsorption lead-ing to loss of enzyme activity, reduced substratespecificity, or altered enthalpy/entropy;2. changes caused by partitioning of the pHenvironment, substrate, products, activators andinhibitors between the enzyme’s microenviron-ment and the bulk soil solution;3. steric hindrance by matrix shielding or occu-pation of active sites by inhibitors; and4. effects of external and internal diffusion.

Immobilization may impact the velocity of phy-tase reactions by causing conformational changesthat render the enzyme denatured or reduce therate of substrate turnover at the enzyme activesite. Likewise, the entropy of immobilized phytasewill be increased due to reduced flexibility, result-ing in an enzyme less likely to achieve conforma-tional requirements for hydrolysis. However, in

some cases immobilization may also reduceenthalpy in comparison to free phytase, resultingin a reduced energy requirement to reach thetransition state (Kelleher et al., 2004). The balancebetween the effects of immobilization on enthalpyand entropy will, at least partially, determine themaximum velocity of the immobilized reaction.

The affinity of an immobilized phytase forits substrate will only be the same as for the freeenzyme when the supporting matrix is eitheruncharged or the ionic strength is great. Whensupports and substrates are similarly charged, asin the case of phytase immobilized on a clay sur-face, the inositol phosphate concentration will bereduced in the microenvironment of the enzyme,and the affinity, reduced. In contrast, if the sup-port and substrate carry opposing charge, as isthe case when phytase is immobilized on metal(hydr)oxides or some organic materials, the inosi-tol phosphate is attracted to the microenviron-ment and the affinity of the reaction mayincrease (Crook et al., 1970; Ladd and Butler,1975). Altered partitioning of inhibitors, activa-tors and products of the catalytic reactionbetween the immobilized phytase and soil solu-tion may also affect the catalytic efficiency ofphytase.

A further consequence of partitioningbetween immobilized phytase and soil solutionwill be an apparent shift in pH optima. This isexplained by accumulation or dissipation ofhydrogen ions in the enzyme’s microenvironment(Violante and Gianfreda, 2000), dependent onthe charge of the supporting surface. These shiftsin pH optima are therefore not due to changes inthe properties of the immobilized enzyme per se,but are an artefact of the difference betweenmeasured pH and that in the microenvironment(Goldstein, 1976). Never the less, it is experimen-tally difficult to separate this potential pH surfaceeffect from well-documented pH-dependentmodifications of conformation or orientation ofthe enzyme on the solid surface (Baron et al.,1999; Quiquampoix 2000; Servagent-Noinvilleet al., 2000).

Reduced activity of enzymes associated withhigh-molecular weight organic substances has, inmost cases, been attributed to steric limitations tothe penetration of substrates to the active site(Goldstein, 1976). It is likely that the affinity ofphytase will vary depending on the soil particle-size fraction the enzyme is associated with (Marx


et al., 2005). Although it has been suggested thatdifferent isozymes, with different affinities, areassociated with various size fractions (Rojo et al.,1990), it is probable that enzymes associated withlarger particle-size fractions have greater accessto substrates than in smaller fractions. Therefore,steric hindrance is generally considered to be themain reason for reduced affinity with decliningfraction size.

Diffusion rates of both phytase and inositolphosphate are clearly important factors thatrestrict the affinity of their reaction in soil.Diffusion of free phytase will depend on its netcharge and size as determined by the amino acidsequence and the degree of modification or sub-stitution (Table 14.1). For example, greater glyco-sylation of fungal phytases compared to bacterialphytases may impact their relative mobility (Leiand Porres, 2003). The mobility of phytase itselfmay not be a major limitation to affinity towardsan unlimited substrate. However, the availabilityof inositol phosphates in soil solution is likely tobe limited (see Celi and Barberis, Chapter 13,this volume) and catalytic reactions will bereduced significantly by either the rate of diffu-sion of substrate to enzyme or vice versa(McLaren and Packer, 1970). Consequently,when diffusion rates are less than the rate ofhydrolysis, a depression in phytase activity maybe observed (Ladd and Butler, 1975; Goldstein,1976; Ciurli et al., 1996). In soil, diffusion rates ofinositol phosphate will depend on the soil type(sorption capacity for inositol phosphate) and alsothe tortuosity of path for diffusion, which in turnis dependent on water content. Importantly, thenumber and distribution of phosphate groups onthe inositol ring will affect the rate of diffusion to,and within, enzyme supports (Engasser andHorvath, 1976) due to the reaction with anionexchange sites. This is likely to affect the speci-ficity of immobilized phytase for the various inos-itol phosphates and stereoisomeric forms.

The kinetics of an enzyme that is embeddedin a porous matrix, such as an organic complex,will be further complicated by rates of internaldiffusion, which will decrease with increasingdepth into the matrix owing to progressive deple-tion of the substrate (Goldstein, 1976). Therefore,the distribution of enzymes in relation to the soilsolid phase will affect the performance of the cat-alytic reaction (Nannipieri and Gianfreda, 1998).For example, phytases located on the external

surface of soil aggregates or organic matrices arelikely to be less affected by diffusional limitationsthan those internal to the structure of the soilaggregate. This has been demonstrated by thefact that the affinity of adsorbed enzymes for sub-strate is much greater when associated withcrushed soil than with intact aggregates (Brahmsand McLaren, 1974).

An important caveat is that when inhibitionby diffusion and other factors, whether chemicalor biological, is concurrent, the combined effect isan apparent amelioration of the initially observedinhibition. For example, if an inhibitor reduces theabsolute activity of adsorbed enzyme and thisreduces the difference between the rate of hydroly-sis and rate of diffusion, diffusional inhibitionof catalytic activity will appear to be increased(Goldstein, 1976). This also has the effect of appar-ently increasing the stability of an immobilizedenzyme. Therefore, in a diffusionally limited sys-tem such as soil, phytase activity may appear toremain stable even though the protein has under-gone considerable denaturation (Goldstein, 1976).

Microbial and proteolytic degradation

Like all enzymes, phytases around plant roots willimmediately encounter a repressive environment,being subject to potential microbial and proteolyticdegradation upon exudation or loss from the cyto-plasm (Tinker and Nye, 2000). Importantly, thereare specific variations in the biochemical nature ofphytases in relation to their susceptibility to prote-olytic degradation, thought to be due to their vul-nerability to conformational change (Simon andIgbasan, 2002). Protein degradation invariablyoccurs at exposed loops on the surface of the mole-cule and directed mutagenesis of A. fumigatus phy-tase has yielded variants that are considerably moreresistant to proteolysis (Fig. 14.4; Wyss et al., 1999b).Phytases from different soil fungi also exhibit varia-tion in their susceptibility to microbial degradation.For example, phytase from P. lycii was more suscep-tible to microbial degradation than that fromA. niger (George et al., 2006). Glycosylation of phy-tases may also further affect their susceptibility tomicrobial and proteolytic degradation and is knownto be highly variable (Wyss et al., 1999b), particu-larly when expressed in heterologous systems. Asdiscussed earlier, protection from microbial and


proteolytic degradation is afforded by immobiliza-tion on to the soil solid phase presumably as a resultof steric hindrance towards degradative agents(Nannipieri et al., 1996; Rao and Gianfreda, 2000;George et al., 2006). Indeed, this stabilization ofimmobilized enzyme most likely contributes to thelevel of endogenous phytase activity that exists insoil. Moreover, it suggests that the soil environmentexerts a selection pressure on the enzyme such thatextended residence differentiates only the mostrobust or most protected proteins.

Inhibition and activation by ionsand metabolites

Most phytases require divalent cations for activ-ity, which is thought to be due to their involve-ment in the active conformation of the phytases(Irving, 1980; Choi et al., 2001). Plant phytasestend to have a broader range of activators thanthose from microorganisms; for example, plantphytases are activated by magnesium, calcium orcobalt ions (Peers, 1953; Nagai and Funahashi,1962; Gibbins and Norris, 1963; Chang, 1966;

Lolas and Markakis, 1977), whereas bacterialphytases are activated solely by calcium ions(Irving and Cosgrove, 1971a; Powar andJagannathan, 1982; Shimuzu, 1992; Kerovuoet al., 1998; Kim et al., 1998a; Choi et al., 2001).In contrast, yeast (Saccharomyces spp.) phytase isactivated by iron(II) and copper(II) ions (Nayiniand Markakis, 1984, 1986), whereas other fungalphytases do not require any specific activation bya cation (Irving and Cosgrove, 1974). It is reason-able to assume that divalent cation activatorssuch as these are abundant in soil and should notlimit the activity of phytases, particularly in thepH range over which most phytases are active.

In contrast, phytase activity can be inhibitedby a wide range of metals, including Ag, Cd, Co,Cr, Cu, Fe, Hg, Mn, Ni, Pb, Sn, W and Zn (Peers,1953; Yamada et al., 1968; Powar andJagannathan, 1982; Nayini and Markakis, 1984;Svenson, 1986; Shimuzu, 1992; Hayes et al., 1999),with mixtures of metal ions having at least additiveeffects on acid phosphatase (Renella et al., 2003).Although the mode of inhibition by metal ions isnot clear, it is suggested that they may competewith activators, cause precipitation of substrates,alter the active conformation of the enzyme orcause steric hindrance of substrate to the active site(Lolas and Markakis, 1977; Gianfreda and Bollag,1996). In particular, metal ions that form insolublesulphides are strong inhibitors (in the order Mn <Co < Cd < Cu < Hg < Ag), which suggests thatinhibition occurs through interaction with sulph-hydryl groups in the active site of the enzyme(Shaw, 1954; Juma and Tabatabai, 1977; Geigeret al., 1998b; Huang and Shindo, 2000b, 2001).Metal ions tend to reduce the velocity but increasethe affinity of the reaction of extracellularenzymes, suggesting that they enhance the bindingof inositol phosphate with the enzyme catalytic site(Huang and Shindo, 2000a,b, 2001). Many ofthese metals occur naturally in soil, albeit at con-centrations that are unlikely to be inhibitory tophytases. However, concentrations of such metalscould be considered inhibitory in some field sitesacutely polluted by human activity.

Inhibition by anions including phosphate(the reaction product), fluoride and arsenate isalso evident and appears to be more potentagainst plant than microbial phytases (Nagai andFunahashi, 1962; Gibbins and Norris, 1963;Chang, 1966; Mandal et al., 1972; Chang andSchwimmer, 1977; Lolas and Markakis, 1977;


90

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00 15 30 45 60 75

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Fig. 14.4. Proteolytic susceptibility of wild-typeand mutated Aspergillus fumigatus phytase.Shown is the activity of A. fumigatus wild-typephytase (●), A. fumigatus S126N phytase mutant(■), A. fumigatus R125L/S126N phytase mutant( ) after incubation in the presence of proteolyticenzymes and A. fumigatus wild-type phytase (▲)after incubation with a proteolytic enzymepreparation that had been pre-treated at 90°C for20 min. (From Wyss et al., 1999b. Reproducedwith permission from the American Society forMicrobiology.)

Hayes et al., 1999). Due to subsequent pleiotropiceffects, intracellular enzymes (most isolated plantphytases) are likely to be more tightly regulated byproduct accumulation than extracellular enzymes(most isolated microbial phytases), which gener-ally show little or no regulation by phosphate(Nannipieri and Gianfreda, 1998). Inhibition byanions can also occur through precipitation ofcationic activators, or by competitive inhibitionwith inositol phosphates for the enzyme active site(Irving and Cosgrove, 1971b). Phosphate is gener-ally in low concentrations in solution and isdepleted rapidly from the rhizosphere, so it isunlikely to inhibit phytase activity. Moreover,product inhibition is generally considered moreeffective against transcriptional regulation of phy-tases than direct inhibition of extracellular phy-tase activity in the soil (Bianchetti and Sartirana,1967; Olander and Vitousek, 2000; Vats andBanerjee, 2004). Notwithstanding this, phosphateconcentrations in soil solution may becomeinhibitory during waterlogging or following appli-cation of manure or mineral fertilizer. Otherpotential inhibitory anions such as arsenate andfluoride are not common in soil solution, whilstanions such as nitrate, sulphate and chloride donot appear to inhibit phosphatases (Juma andTabatabai, 1977).

In contrast, inhibition of microbial phytasesby chelating agents such as citrate, oxalate, tartrateand ethylenediaminetetraacetate (EDTA) appearto be more acute than that of plant phytases (Irvingand Cosgrove, 1971a; Nayini and Markakis, 1984;Shimuzu, 1992; Yoon et al., 1996; Kerovuo et al.,1998; Choi et al., 2001). Chelating agents couldinhibit enzyme activity by binding with activators,although the effect may be mitigated through com-plexation with metals that would be inhibitory tophytases. The concentration of plant and microbialmetabolites, such as citrate, oxalate and malate, isgreatly enhanced within the rhizosphere (Tinkerand Nye, 2000) and could therefore have a majoreffect on phytases within this zone.

Denaturation by soil environmentalfactors

The pH environment in soil is likely to be moreextreme and temporally variable than that of thecytoplasm. Despite this, the biological range of

phytases (Table 14.1) appears well able to copewith this environment, having optimal pH forcatalytic reactions ranging from 2.2 to 8.6(Irving, 1980; Nayini and Markakis, 1986).However, most discrete plant and microbial phy-tases have narrow single optima in the range ofpH 3.5–7.5, and show significant declines in cat-alytic activity with small changes in pH on eitherside of this optimum (Peers, 1953; Nagai andFunahashi, 1962; Gibbins and Norris, 1963;Chang, 1966; Irving and Cosgrove, 1971a;Chang and Schwimmer, 1977; Lolas andMarkakis, 1977; Basha, 1984; Nayini andMarkakis, 1984; Greiner et al., 1993; Yoon et al.,1996; Kerovuo et al., 1998; Kim et al., 1998a;Hayes et al., 1999; Liu et al., 1999; Choi et al.,2001; Quan et al., 2004). Soil environments areunlikely to be conducive to optimal catalyticactivity of phytases, as the pH is unlikely to beeither optimal or remain stable at this optimalvalue. Notwithstanding this, phytase activity iso-lated directly from soil tends to have a broaderrange of pH optima (Svenson, 1986), and discretephytases have been shown to have multiple andbroad pH optima (Irving and Cosgrove, 1974;Greiner et al., 1993; Casey and Walsh, 2003;Brugger et al., 2004; Chadha et al., 2004;Dharmsthiti et al., 2005). Moreover, it is also nowpossible for phytases to be specifically engineeredfor broader pH optima (Mullaney et al., 2002;Tomschy et al., 2002). Although soil pH isunlikely to be optimal for phytase activity, it isalso unlikely to lead to complete denaturation ofthe phytase protein, the structures of which arestable (activity was recoverable) against pH envi-ronments ranging from pH 1.2 to 11 (Yamadaet al., 1968; Shimuzu, 1992).

Phytases have generally been found to betemperature-stable with optimum activity in therange of 45–57ºC (Table 14.1). Beyond this, phy-tase activity tends to decline due to thermaldenaturation with total denaturation occurring at~80ºC (Irving, 1980). Importantly, some phytases(e.g. from A. fumigatus) are capable of re-formingtheir active conformation following exposure tohigh temperature (Wyss et al., 1998). Differencesin glycosylation of phytases may also affect ther-mostability (Han et al., 1999), and specific modifi-cations to the amino-acid sequence have beenshown to increase tolerance to extreme tempera-ture (Lehmann et al., 2000, 2002). Temperatureis unlikely to denature phytases in soil under


normal conditions, although exposed surface lay-ers will approach or exceed denaturation temper-atures in certain locations and certain times ofthe year. Total loss of phytase activity by thermaldenaturation may also occur in soil during burn-ing of above-ground vegetation (Saa et al., 1993;Staddon et al., 1998). This has implications forthe recovery of biological cycling of inositol phos-phate following natural forest fires or those usedin agricultural and ecological management.Importantly, interaction with soil constituents canalso make phytases less prone to thermal denatu-ration. For example, phytase isolated from mungbean was shown to be less sensitive to tempera-ture denaturation when associated with inositolphosphates and divalent cations such as calcium(Mandal et al., 1972). Such interactions areassumed to alter the conformation of the enzyme,making it less susceptible to denaturation (Kimet al., 1998b; Choi et al., 2001).

At typical soil temperatures standard thermo-dynamic principles will apply, such that enzymekinetics will be temperature-dependent, with ratesincreasing up to the range of optimal temperaturesmentioned above. Freezing may reduce the activ-ity of extracellular phytases through denaturation(Pettit et al., 1977) and may affect the rates ofenzyme reactions by changing the ionization of allreactants and the conformation of the protein(McLaren and Packer, 1970). The severity offreezing conditions is also important. Slow freezinggenerally leads to localized concentrations of reac-tants and an increase in the affinity of the system,whereas during rapid freezing the reactantsremain homogenized and activity is severelyretarded (McLaren and Packer, 1970).

Water is essential for phytase activity, beingthe medium in which reactions occur. In general,enzyme activities tend to decline with drying(Gianfreda and Bollag, 1996). As soil dries, denat-uration of extracellular enzymes occurs (Rao et al.,2003) due to unfolding of secondary structures.However, some enzymes regain their activity withrehydration, although self-association hinders thecomplete recovery of tertiary structures (Noinvilleet al., 2004). Although small changes in waterpotential may lead to modified protein structureand enzyme activity (Reyes et al., 2005), hydrolyticenzymes actually require very little water to beactive. For example, extracellular urease requires1.3 moles of water per mole of side chain polargroups to be active, suggesting that enzyme activi-

ties, including phytase, may remain functionaleven in air-dried soil (McLaren and Packer, 1970).Interestingly, plant, microbial and fungalhydrophyllins (hydrophilic extracellular proteins)can reduce conformational changes to extracellu-lar enzymes usually observed with reduced waterpotential, and thus avoid the loss of enzyme activ-ity with drying (Reyes et al., 2005).

A secondary effect of soil drying is anincrease in ionic strength in soil solution. This mayincrease passive loss of phytases from microorgan-isms through osmotic stress and increase phytaseactivity in solution by reducing the adsorption ofphytase through neutralization of electrostaticforces. Inhibitory or activatory effects of soil solu-tion salts will also be concentrated by drying(Gianfreda and Bollag, 1996). In addition to dry-ness, excess water may impact phytase activity insoil. Waterlogging tends to limit extracellularenzyme activity (Freeman et al., 1996; Gianfredaand Bollag, 1996; Kang and Freeman, 1999;Chacon et al., 2005), through inhibition by metalions such as iron and manganese in the reducedstate, which are more soluble than their oxidizedequivalents (Pulford and Tabatabai, 1988). As withmetal ion toxicity, this effect appears to be miti-gated by immobilization of extracellular enzymeson solid surfaces (Goel et al., 1998).

Finally, phosphatases can be degraded bylight, particularly short-wave radiation, such asultraviolet-B (Espeland and Wetzel, 2001).This will be of little consequence to phytase in soil,except when enzymes are exposed to light either atthe soil surface or following tillage (Nannipieri and Gianfreda, 1998). Photodegradation may bemore important when phytases move from soilto aquatic environments. Despite the inherentlonger exposure to light radiation following thistransition, movement through the environment asorganic–enzyme complexes will afford phytasessome protection from photodegradation (Wetzel,1992, 1993; Espeland and Wetzel, 2001) as willcomplexation with clay (Tietjen and Wetzel, 2003).

Concluding Remarks and FutureDirection

It is evident that the soil environment has a majoreffect on the ability of phytases to hydrolyse phos-phorus from inositol phosphates. Phytases released


to soil from plants and microorganisms, or thoseadded to soil in manures from animals fed modi-fied diets, will operate under suboptimal condi-tions. Major factors affecting phytase activity insoil include susceptibility to microbial and prote-olytic degradation, variability of both pH andmoisture content and, importantly, the immobi-lization and denaturation of phytase by the soilsolid phase. Given these factors and that inositolphosphates react strongly in soil through precipita-tion and adsorption reactions, which can signifi-cantly affect their availability to phytases (see Celiand Barberis, Chapter 13, this volume), it is per-haps not surprising that inositol phosphates appearto resist mineralization and form a major con-stituent of the organic phosphorus in most soils.However, until we understand the role of the soilphytase activity that is retained in the long termand quantify the actual biological cycling of inosi-tol phosphates in soil, it is difficult to speculate onthe true importance of extracellular phytases in thesoil environment. It remains to be proven whethersuch baseline phytase activity has an integral eco-logical role, such as warning microorganisms intheir niche of temporal changes in the presence ofinositol phosphates, as suggested for other phos-phatases in Burn’s hypothesis, or whether theseactivities are simply fortuitously retained due totheir protection against microbial degradation byadsorption to soil particles. However, the apparentinterspecific variation in phytases and differencesbetween functional groups (intracellular vs. extra-cellular) suggest that some advantage is gained byproducing phytases with greater longevity inthe soil environment, and that accidental retentionof phytase activity would be a less-favoured conclusion.

The challenge for future research is to betterunderstand the efficiency of phytase–inositolphosphate interactions in soil. In particular, thefollowing aspects require investigation:

1. the factors that control the availability of inos-itol phosphates for interaction with phytases;2. the importance of the differing biochemicaland physiological properties of phytases fromvarious biological sources (e.g. bacteria, fungi,plants) and different classes of enzyme (e.g. histi-dine acid phosphatases vs. β-propeller phytase vs.purple acid phosphatases; see Mullaney andUllah, Chapter 7, this volume) on the dephos-phorylation of inositol phosphates in soil;

3. the role of phytases in the dephosphorylationof the range of inositol phosphates, includingphosphorylated inositol stereoisomers, found innature; and4. the role of phytases in ecosystem function andtheir significance for the turnover of inositolphosphates as a component of the soil phospho-rus cycle.

Importantly, there is opportunity to exploit thenatural variability in the biochemical characteris-tics of phytases or that which can be generatedthrough protein engineering. Interspecific differ-ences in the susceptibility of phytases to microbialdegradation are evident, and the capacity to gen-erate phytases that are less prone to proteolyticdegradation is increasing. Similarly, phytaseshave been identified that are active over a rangeof soil pH and again it is possible that phytaseswith a broader range of pH optima can be engi-neered. Genetic variability in the susceptibilityof phytases to immobilization by adsorptionand subsequent degradation is also evident.Collectively, manipulation of these biochemicalcharacteristics may make it possible to tailor spe-cific phytases for optimal function in a range ofsoil environments and thus more effectively man-age the interaction between phytases and inositolphosphates.

The question remains, however, whethersuch changes would have any impact on thekinetics of phytase reactions in soil and thus thebioavailability of inositol phosphates. To addressthis it must be established whether soil–plant sys-tems are already ‘optimized’ with respect to phy-tase activity and function at a ‘natural’ capacity,whereby the presence and/or accumulation ofinositol phosphates over the long term isinevitable. Importantly, the ecological signifi-cance of phytase and inositol phosphates insoil–plant systems must be determined.

We now have a range of experimental toolsand analytical procedures that allow us to morethoroughly address some of these questions. Forexample, transgenic plants that express heterolo-gous phytases can act not only as a delivery sys-tem for specific phytases to the rhizosphere, butalso as bio-indicators to determine whether phy-tases with specific biochemical traits are effectiveat improving the bioavailability of inositol phos-phates. This is important with regard to the utilization of inositol phosphates that are either



endogenous to soil or added through animalmanure and plant residues.

At present, we are well poised to address keyknowledge gaps in understanding critical param-eters that control the turnover of inositol phos-phates in soil. This will not only contribute to our

understanding of the soil phosphorus cycle inboth natural and agricultural ecosytems, but mayalso provide opportunity to improve phosphorusefficiency in agriculture, reducing the reliance onphosphorus fertilizer and any consequent envi-ronmental degradation.

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15 Plant Utilization of InositolPhosphates

Alan E. Richardson1, Timothy S. George2, Iver Jakobsen3 andRichard J. Simpson1

1CSIRO Plant Industry, PO Box 1600, Canberra, ACT 2601, Australia;2Scottish Crop Research Institute, Invergowrie, Dundee DD2 5DA, United Kingdom;

3Risø National Laboratory, Biosystems Department, Roskilde, DK 4000, Denmark

Inositol phosphates are a major component ofthe organic phosphorus in most soils, yet ourunderstanding of the availability of inositol phos-phates to plants is limited. Phosphorus deficiencyis a major constraint to the growth of plants inmany agricultural systems throughout the worldand influences species composition in naturalecosystems (e.g. Wassen et al., 2005). This isdespite the fact that soils generally contain a rela-tively large amount of total phosphorus. Thistotal phosphorus includes organic and inorganicforms that are bound to soil particles by adsorp-tion reactions or are present in mineral and pre-cipitated complexes. Agricultural productionsystems are therefore reliant on the application ofphosphorus-based fertilizers to meet the phos-phorus requirement of plants. Such fertilizers arecomposed primarily of soluble inorganic phos-phate processed from rock phosphates, or arederived from animal manures and other biologicalresidues. However, much of the inorganic phos-phate that is added to soil is rapidly ‘fixed’ (byadsorption and precipitation reactions) or isimmobilized into organic phosphorus by soilmicroorganisms (Sanyal and De Datta, 1991;Oberson and Joner, 2005; Pierzynski et al., 2005),with the result that only a relatively small propor-tion of the phosphorus applied as fertilizer istaken up by plants. Consequently, there is theneed to better understand how plants acquirephosphate from ‘endogenous’ forms of soil phos-

phorus, from applied sources of organic phospho-rus, or from phosphorus that accumulates underdifferent management systems.

Accumulation of organic phosphorus and itsutilization by plants are of particular interest,because organic phosphorus accounts for at least50% and up to 80% of the total phosphorus inmany soils (Harrison, 1987). Whilst much of theorganic phosphorus in soil is associated withhigh-molecular weight fractions, a large partcomprises phosphate monoesters. Of this, variousstereoisomers of inositol penta- and hexakisphos-phates are the major constituents and account forapproximately 50% of the total organic phospho-rus (Anderson, 1980; Turner et al., 2002b; seeTurner, Chapter 12, this volume). Inositol phos-phates are also the major storage compound forphosphorus in plant seeds, in which salts of myo-inositol hexakisphosphate (phytate) account for~70% of the total seed phosphorus (see Raboy,Chapter 8, this volume). Inositol phosphates (pri-marily as phytate) are thus a significant compo-nent of the dietary phosphorus intake of animalsin intensive livestock industries. For monogastricanimals in particular (i.e. swine and poultry) inos-itol phosphates in manures may therefore beimportant to the phosphorus cycle in soil–plantsystems fertilized with manure (see Leytem andMaguire, Chapter 10, this volume). However, wehave little understanding of the reactions of inosi-tol phosphates in soil or their ‘biological

availability’ to plants and microorganisms in soilenvironments.

This chapter considers the importance of inos-itol phosphates as a potential source of phosphorusfor plant nutrition. In particular, the activity of phy-tase in the rhizosphere and its contribution to thehydrolysis of inositol phosphates in soil is consid-ered. Better knowledge of the role of inositol phos-phates in the phosphorus nutrition of plants mayimprove our understanding of ecosystem function,and may also provide strategies to improve the effi-ciency of phosphorus-fertilizer use in different agri-cultural systems throughout the world.

Phosphorus Nutrition of Plantsand Adjustment to Phosphorus

Deficiency

Organic forms of phosphorus are not directlyavailable to plants, which take up phosphorus assoluble phosphate anions (HPO4

2− and H2PO4−).

This occurs primarily across the plasma mem-brane of the root epidermis, through root hair cellsor by mycorrhizae that are associated with roots.In most soils the concentration of phosphate in soilsolution is low. Phosphorus deficiency occurs whenthe capacity for replenishment of phosphate orrates of diffusion are insufficient to meet plantrequirements (Bieleski, 1973; Seeling and Zasoski,1993). Plants have evolved a range of mechanismsthat improve their capacity to acquire phosphatefrom the external environment and to maximizeinternal phosphorus utilization when deficient.These mechanisms are reviewed in detail else-where (e.g. Raghothama, 1999, 2005; Vance et al.,2003) and in summary include:

1. morphological changes to root structure suchas rate of root growth, increased total and specificroot length, the degree of root branching and theabundance and length of root hairs (Lynch, 2005;Hill et al., 2006), all of which allow plants toexplore greater volumes of soil;2. association with soil microorganisms, in par-ticular mycorrhizal fungi and non-symbioticmicroorganisms that can enhance either theavailability or the uptake of phosphate from soil(Richardson, 2001; Jakobsen et al., 2005); and3. biochemical processes that occur at theroot–soil interface and within the rhizosphere.This includes induced expression of specific

proteins for the transport of phosphate across theplasma membrane of root cells or those whichfacilitate its transfer from mycorrhizal fungi(Rausch and Bucher, 2002), modification to rhi-zosphere pH (Hinsinger, 2001), the release ofroot exudates (e.g. low-molecular weight organicanions) that improve phosphorus availabilitythrough increased solubilization of both inor-ganic and organic phosphorus pools (Hocking,2001; Ryan et al., 2001) and the release of phos-phatase enzymes (in particular acid phospho-monoesterases and -diesterases), which arerequired for the hydrolysis of organic phosphorussubstrates (Richardson et al., 2005).

Many plant species form symbiotic associationswith mycorrhizal fungi, with the association gener-ally being characterized by a mutualistic exchangeof carbon from the plant in return for mineralnutrients from the soil, primarily phosphate(Smith and Read, 1997). Of particular note arethe ectomycorrhizal fungi, which form associ-ations predominantly with woody plants, and thearbuscular mycorrhizae, which associate with themajority of agricultural species. Characteristic ofectomycorrhizal infections is the formation ofmycelial sheaths that envelop plant roots withhyphae that, although associated with the cellwall, are external to root cells. Arbuscular mycor-rhizae have inter- and intracellular hyphae thatpenetrate the wall and plasma membrane of rootcortical cells with the formation of haustoria-likearbuscules within plant cells. In both instances, itis well established that the fungal mycelia/hyphae increase significantly the surface area ofplant roots and provide greater contact with soilallowing enhanced uptake of phosphate (Jakobsenet al., 2005). There is little evidence to indicatethat mycorrhizal fungi have access to pools of soilphosphorus other than those available to plants(Bolan, 1991; Joner et al., 2000), although it hasbeen suggested that phosphatase activity in myc-orrhizae may provide plants with increasedaccess to soil organic phosphorus (Tarafdar andMarschner, 1994b; Feng et al., 2002).

Phosphatases and the utilization of soilorganic phosphorus

Organic phosphorus in soil and soil solution isnot directly available to plants and must first be

Plant Utilization of Inositol Phosphates 243

hydrolysed by phosphatase enzymes to releasethe phosphate anion. Dephosphorylation mayoccur either in the external root environment or, ifsubstrates are soluble and able to diffuse throughthe root apoplast, within the cell wall space or atthe root endodermis. Plants produce a range ofextracellular acid phosphatases that are associatedwith either root cell walls (i.e. the root apoplasm)or released into the external environment as freeenzymes (e.g. Dracup et al., 1984; Barrett-Lennardet al., 1993; Tadano et al., 1993; Li et al., 1997;Gilbert et al., 1999; Hayes et al., 1999; Zhangand McManus, 2000). Characterization of pur-ple acid phosphatase genes (see Mullaney andUllah, Chapter 7, this volume) from arabidopsis(Arabidopsis thaliana (L.) Heynh.) and white lupin(Lupinus albus L.) has confirmed the extracellularnature of these enzymes and their inducedexpression in response to phosphorus deficiency(Haran et al., 2000; Wasaki et al., 2000; Milleret al., 2001). Acid phosphatase activities have like-wise been reported in isolated mycelium of myc-orrhizal fungi. However, as with plant roots,cell-bound phosphatase activity generally domi-nates over extracellular forms (Straker andMitchell, 1986; Kroehler et al., 1988; Ho, 1989;Antibus et al., 1992; McElhinney and Mitchell,1993; Joner and Johansen, 2000, Olsson et al.,2002). Colonization of roots by both ectomycor-rhizal and arbuscular mycorrhizal fungi has alsobeen shown to increase the acid phosphataseactivity of roots (Antibus et al., 1981; Dodd et al.,1987; MacFall et al., 1991; Ezawa and Yoshida,1994; Fries et al., 1998), although its contributionto the extracellular component of root activity isunclear (Ezawa et al., 2005). Presence of extracellu-lar acid phosphatase activity in hyphae associatedwith roots has been visualized histochemically forboth arbuscular and ectomycorrhizae (Feng et al.,2002; Alvarez et al., 2005) and for intact plantroots (Dinkelaker and Marschner, 1992; Griersonand Comerford, 2000).

Release of extracellular phosphatases fromplant roots is also consistent with greater activityin the rhizosphere of soil-grown plants, particu-larly in response to phosphorus-deficient conditions(reviewed by Richardson et al., 2005). Greaterphosphatase activity is generally accompanied bya depletion of soil organic phosphorus from therhizosphere (Tarafdar and Jungk, 1987; Chenet al., 2002; George et al., 2002; Liu et al., 2004).Greater activity of acid phosphatases has simi-

larly been correlated with hyphal length of ecto-mycorrhizae associated with roots (Häussling andMarschner, 1989) and in some cases with themycelial density of both arbuscular and ectomyc-orrhizae in root-free soil compartments (Tarafdarand Marschner, 1995; Feng et al., 2002; Liu et al.,2005), although this has not been observed in allcases ( Joner and Jakobsen, 1995; Joner et al.,1995). In the study by Liu et al. (2005) using radi-ata pine (Pinus radiata D. Don.), soil phosphataseactivity was positively correlated with the lengthdensity of mycelium in root-free zones of soil andwas associated with a significant depletion of soilorganic phosphorus.

However, the relative importance of phos-phatases produced by plant roots, mycorrhizae orother free-living microorganisms in the rhizos-phere, the activity and numbers of which are alsosubstantially larger around roots ( Jakobsen et al.,2005), is not well understood. Whilst it is evidentthat phosphatases in the rhizosphere are effectivefor the depletion of organic phosphorus in variousoperationally defined pools (e.g. extractable insodium bicarbonate, sodium hydroxide), there is aneed to investigate the interaction of specific acidphosphatases and the utilization of defined organicphosphorus substrates (Richardson et al., 2005).

Implications for the utilization of inositolphosphates by plants in soil

Despite the abundance of inositol phosphates insoil, their use by plants will depend on variousfactors that include:

1. The proximity to roots of inositol phosphates in soil.Roots (with or without mycorrhizae) must effec-tively explore soil to interact with substrate andto capture phosphate released by hydrolysis incompetition with other reactions of phosphate insoil (e.g. immobilization by soil microorganismsor physical and chemical fixation). Roots andmycorrhizae are potentially well suited to exploitphosphorus in patchy environments such asorganic layers, where hyphal proliferation andpenetration into soil pores may be stimulated(Ravnskov et al., 1999; Gavito and Olsson, 2003;Hodge, 2004).2. The solubility and mobility of inositol phosphateseither within the soil solution or the root apoplasm.Soil solution contains a wide range of organic

244 A.E. Richardson et al.

phosphorus compounds (Wild and Oke, 1966;Martin, 1970), and phosphate monoesters,including inositol hexakisphosphates, have beenidentified as a component of soil leachate(Espinosa et al., 1999; Toor et al., 2003).However, their concentration in soil solution islikely to be small. Moreover, due to their highcharge density, inositol phosphates are notexpected to diffuse freely in soil solution andwithin plant cell walls. Inositol phosphates adsorbstrongly to clays, metal oxides and organic matterwith sorption capacities being equivalent to about4 times that of phosphate anions on a per mole-cule basis (Anderson et al., 1974; Shang et al.,1992; Celi and Barberis, 2005; see Celi andBarberis, Chapter 13, this volume). Dependingon pH, inositol phosphates also form sparinglysoluble precipitates with a range of cations, withcalcium and magnesium complexes beingpredominant under alkaline conditions andaluminium and iron complexes under acidicconditions (Jackman and Black, 1951). Thesereactions contribute to the stabilization of inositolphosphates in soil and are major factors that willaffect its concentration in soil solution, mobilitywithin the solution phase and susceptibility toenzyme hydrolysis (Tang et al., 2006).3. The presence and activity of phytases of either plant ormicrobial origin, their location in or around plant rootsand their capacity to effectively interact with substrate.At present, we have limited knowledge of theinteraction of phytase with inositol phosphatesin the soil–root environment and poor under-standing of the rate-limiting steps for substratedephosphorylation (see George et al., Chapter 14,this volume).

Utilization of Inositol Phosphatesby Plants Grown in Axenic Culture

Plants have limited capacity to access phosphorusfrom inositol phosphates relative to other organicsubstrates when grown under controlled conditionswhere availability of substrate is not expected tobe limited (Hayes et al., 2000b; Richardson et al.,2001b). Using a number of grass and pasturelegumes, Hayes et al. (2000b) showed that plantsgrown in sterile agar were unable to effectivelyobtain phosphorus from myo-inositol hexakisphos-phate (supplied as sodium-phytate), even when

provided at concentrations that were up to 20-fold higher than the phosphate concentrationrequired for the maximum growth of each species.Similarly, wheat (Triticum aestivum L.) had limitedability to acquire phosphorus from sodium-phytatewhen compared to a range of other monoesterand diester substrates. Ester-bonded phosphatesother than phytate produced equivalent growthand phosphorus uptake as plants supplied withinorganic phosphate (Richardson et al., 2000). Inboth cases the inability to utilize phosphorusfrom phytate was considered to be associated withlow levels of extracellular phytase that was releasedfrom roots into the external medium. This wasconfirmed by significantly improved plant growthand phosphorus uptake of subterranean cloverseedlings when a purified phytase was added tothe growth media (Hayes et al., 2000b).

Inoculation of plants with microorganismsthat possess phytase activity also improved thephosphorus nutrition of plants supplied withphytate. Phosphorus uptake by wheat, subter-ranean clover and a range of other plant specieswas significantly greater in the presence of anisolate of Pseudomonas sp. that was selected forextracellular phytase activity (Richardson andHadobas, 1997; Richardson et al., 2000, 2001a).Growth promotion that is attributable to thephytase activity of bacteria has similarly beenreported for plants inoculated with Bacillus amy-loliquefaciens and a range of Burkholderia spp.(Idriss et al., 2002; Unno et al., 2005). Collectively,these studies indicate that plant roots do not pos-sess an extracellular phytase activity that is effec-tive for the utilization of phosphorus frominositol phosphates and that this inability can becomplemented by the phytase activity ofmicroorganisms.

Phytase activity of plant roots

Phytase activity of roots has been measured for arange of plant species and has generally beenshown to be absent or to constitute a small com-ponent only of the extracellular phosphataseactivity of plant roots (Barrett-Lennard et al.,1993; Asmar, 1997; Bosse and Köck, 1998;Gilbert et al., 1999; Hayes et al., 1999; Richardsonet al., 2000; Lung and Lim, 2006). In wheatseedlings, for example, phytase accounted for


between 1% and 5% of the total phosphomo-noesterase activity of root extracts (as determinedusing para-nitrophenyl phosphate as a model sub-strate), and although evident in whole root assays,was not detected as an extracellular activity inroot exudates (Richardson et al., 2000). This wasevident irrespective of plant phosphorus statusand despite a twofold increase in the level of totalphosphomonoesterase activity in exudates inresponse to phosphorus deficiency. Likewise, intobacco roots (Nicotiana tabacum L.), phytase con-stituted 2.7% and 4.6% of the total activity inphosphorus-sufficient and -deficient plants,respectively, and although detectable in the exter-nal medium over 14 days, was not present as asignificant root-released extracellular enzyme(Lung and Lim, 2006). Phytase activities were,however, present in both whole root extracts androot-associated cell wall fractions. A cell wall com-ponent of extracellular phytase has similarly beenobserved for subterranean clover without measur-able release of enzyme activity (Hayes et al., 1999;George et al., 2004). White lupin also releases acidphosphatase from roots in response to phosphorusdeficiency, but without significant release of phy-tase activity (Gilbert et al., 1999). The exception tothese observations is the phytase activity in rootexudates from a range of plant species reportedby Li et al. (1997), where activities were substantialand equivalent to the total acid phosphatase.

Lack of an exuded phytase activity fromplant roots is consistent with observations of thein situ localization of phytase activity in maize(Zea mays L.) roots, where it was predominantlyconfined to the root endodermis (Hübel andBeck, 1996). Subsequently, two phytase genes(with similarity to histidine acid phosphatases)were cloned from maize and, while expression ofthese genes is consistent with their role in themobilization of phytate in seeds, one of the geneswas also expressed in the endodermis, pericycleand rhizodermis of mature roots (Maugenestet al., 1999). Analysis of this gene and protein,however, provided no direct evidence for itsrelease as an extracellular enzyme. It was hypoth-esized that the role of the phytase was in themobilization of endogenous phytate in plantroots, such as those deposited as phosphorus-richgloboids in pericycle and endodermis cells(Campbell et al., 1991; Van Steveninck et al., 1994;Hübel and Beck, 1996). More recently, Xiao et al.(2005) identified an extracellular phytase (MtPhy1)

from barrel medic (Medicago truncatula Gaertn.),which was shown to be secreted to the root cellwall. Whether this phytase is also released intothe external soil environment remains to bedetermined, as does its effectiveness in allowingbarrel medic to utilize inositol phosphates. Themedic phytase is a purple acid phosphatase thathas similarity to a phytase identified in soybean(Glycine max L. Merr.), which, from its pattern ofexpression in cotyledons, was considered to beinvolved in the mobilization of phytate duringseed germination (Hegeman and Grabau, 2001).Nonetheless, the identification of an extracellularphytase in medic roots provides new opportunityfor furthering our understanding of the functionalsignificance of phytases in plant roots.

Phytase genes from different sources havebeen expressed in plants to facilitate the under-standing of their role in the utilization of inositolphosphates. Richardson et al. (2001a) showed thatexpression of the phyA gene from Aspergillus nigerin arabidopsis improved the growth and phos-phorus nutrition of plants supplied with sodium-phytate. This ability was associated with therelease of phytase as an extracellular enzymefrom the roots of the transgenic plants. In com-parison, wild-type plants or control plants (whichalso expressed phytase but without a signal pep-tide for extracellular targeting of the enzyme) didnot respond when supplied with phytate(Richardson et al., 2001a). Enhanced phosphorusnutrition of transgenic plants that release PhyA tothe rhizosphere has since been demonstrated fortobacco and subterranean clover (George et al.,2004, 2005c). A 70-fold increase in the activity ofexuded phytase resulted in significantly improvedability of the plants to acquire phosphorus fromphytate (Fig. 15.1).

The effectiveness of extracellular release ofheterologous phytases in plants has similarlybeen demonstrated by expression of a consensus(fungal) phytase in transgenic potato (Solanumtuberosum L.; Zimmermann et al., 2003), theβ-propeller phytase from B. subtilis (168phyA) inboth tobacco and arabidopsis (Lung et al., 2005)and, more recently, the expression of the medicMtPhy1 phytase in arabidopsis (Xiao et al., 2005).In a number of cases, these phytases have beenshown to be equally effective when expressed inplants either with constitutive promoters (i.e.the CaMV35S promoter) or with promotersderived from phosphate transport genes (e.g. the



promoter from the AtPht1,2 gene from arabidop-sis), which direct gene expression predominantlyto root hair cells and are induced under condi-tions of phosphorus deficiency (Mudge et al.,2003; Zimmermann et al., 2003; Xiao et al., 2005).These observations are significant, because theyprovide further evidence that plants do not havean innate ability to utilize phosphorus from inosi-tol phosphates and, in soil environments, may bedependent on microbial-mediated mineralization.

Growth and Phosphorus Nutritionof Plants in Soil with Exogenous

Substrate

The ability of plants to utilize phosphorus frominositol phosphates has been of long-standing inter-est in plant nutrition (e.g. experiments by Rogerset al., 1940), and a number of key studies have

investigated their effectiveness compared to inor-ganic phosphorus for plants grown in sand or soil(Martin, 1973; Tarafdar and Claassen, 1988; Becket al., 1989; Adams and Pate, 1992; Findenegg andNelemans, 1993). These studies have generallyshown that phosphate from myo-inositol hexak-isphosphate is available to plants, but its availabil-ity is dependent on the level of substrate supplyand the phosphorus-sorption characteristics of thegrowth medium. In quartz sand, with low phos-phorus-sorption capacity, myo-inositol hexakisphos-phate (supplied as sodium- or calcium-phytate) wasequally available as inorganic phosphate to lupins(L. albus and L. angustifolius L.) when supplied adlibitum at 0.5 mM (Adams and Pate, 1992), andsimilarly was available to maize when applied atgreater concentrations (10 mM and above), wherethe total amount of phosphorus supply was well inexcess of plant requirements (Findenegg andNelemans, 1993). However, when supplied at alower rate (0.2 mM, and at a total phosphorus

Trifoliumsubterraneum

Shoot dry weight(mg/plant)

28.1 40.7 51.8 47.9

Shoot phosphorus(µg P/shoot)

48.3 103.5 299.0 305.3

Exuded rootphytase activitya

(nKat/g root dry wt)

− 1.3 107.9 −

aActivity for wild-type plants was 0.6 nKat/g root dry weight.

ex::phyA ex::phyANull segregantex::phyA

No Pmyo -Inositol hexakisphosphate

(sodium-phytate)Na2HPO4

Fig. 15.1. Growth and phosphorus nutrition and activity of phytase exuded from the roots of transgenicTrifolium subterraneum. Shown are plants that release the Aspergillus niger phytase (ex::phyA) as anextracellular enzyme and its corresponding null segregant transgenic control line. Plants were grown for28 days in sterile agar either without added phosphorus (No P) or with phosphorus supplied as sodium-phytate (myo-inositol hexakisphosphate) or disodium phosphate (Na2HPO4) at 0.8 mM (with respect tophosphate). (From George et al., 2004, in which experimental details are reported.)


supply that was about 3 times that of the plantsuptake requirements), calcium-phytate was ineffec-tive relative to inorganic phosphate. Significantly,the phosphorus nutrition of maize plants grownunder these conditions was improved when phy-tase was added to the sand medium, indicatingthat rate of hydrolysis, as well as availability of sub-strate, was important (Findenegg and Nelemans,1993). Hydrolysis of phytate was also shown tooccur in the sand at high rates of substrate supply,both without the addition of phytase and in theabsence of plants, which indicated that microbialmineralization occurred.

Inositol phosphates are less effective for thephosphorus nutrition of plants when added tosoils. Whilst some studies have shown them to beequivalent to either other organic phosphorussubstrates or to inorganic phosphate (Tarafdarand Claassen, 1988), it has generally been shownthat plant acquisition of phosphorus from exoge-nously added phytate is poor, and essentially afunction of the phosphate-sorption characteristicsof the soil (Beck et al., 1989; Findenegg andNelemans, 1993). This is shown in Fig. 15.2,where the growth of wild-type tobacco plants intwo soils of differing phosphate-sorption capacitywas restricted compared with that for equivalentrates of inorganic phosphate, with greater restric-tion occurring in the higher phosphorus-fixingsoil (A. Richardson, 2006, unpublished data).This is consistent with observations that, com-pared with RNA, glycerophosphate and inor-ganic phosphate, phytate was poorly available tolupin plants in a low phosphorus soil (Adams andPate, 1992) and that phytate was essentially notavailable to maize plants in three contrasting soilswhen supplied at different rates (Findenegg andNelemans, 1993). The limited capacity of plantsto access phosphorus from exogenously suppliedphytate has also been demonstrated using radio-actively labelled substrate. Martin and Cartwright(1971) showed no evidence for plant uptake of 32Pfrom labelled myo-inositol hexakisphosphate byryegrass (Lolium perenne L.) when supplied at a rateof either 20 or 200 mg P/kg soil in two highphosphorus-fixing soils. Substantial uptake occurredin low phosphorus-fixing sand, but only at thehigher rate of phytate supply.

Soil microorganisms may be important forplant access to inositol phosphates in soil irrespec-tive of the fact that its reactivity (i.e., adsorptionand precipitation) is a major factor determining

availability to plants. This is particularly so giventhat plants do not possess high intrinsic phytaseactivity, yet in many cases are able to utilize phos-phorus from phytate when supplied at high rates(Fig. 15.2). Martin (1973) investigated the supplyof labelled myo-inositol hexakisphosphate to wheatplants grown in soils that were either sterilized orre-inoculated with a mixed population of rhizos-phere bacteria or specific isolates that possessedphytase activity. Uptake of 32P by the plants wasessentially dependent on the rate of substrate sup-ply, and no major differences were observed inthe amount of 32P that was taken up by the plantsin the various soil treatments. However, the radio-label was incorporated into the soil microbial bio-mass in all soils and significant mineralizationoccurred through time. On the contrary, Hübeland Beck (1993) found no evidence for depletionof labelled phytate in the rhizosphere of maize.

Improved phosphorus nutrition of a range ofplant species supplied with phytate in a phospho-rus-fixing sand-vermiculite medium has beenobserved after inoculation with soil micro-organisms (Richardson et al., 2001b). Findeneggand Nelemans (1993) also showed that the avail-ability of phosphorus from phytate to maize inthree soils of differing phosphorus-sorption capac-ity was improved by the addition of phytaseenzyme, albeit at rates and substrate concentra-tions that were up to tenfold greater than thatrequired for plants grown in sand. More recently,using transgenic tobacco plants that release PhyA,George et al. (2005c) showed that plant phospho-rus nutrition was increased by up to 52%, com-pared with a wild-type and a transgenic control,in two soils supplied with calcium-phytate.Absolute growth of these plants was, however, stillsignificantly less than that for plants that receivedan equivalent amount of inorganic phosphate.

Significance of mycorrhizae for utilizationof inositol phosphates

The contribution of mycorrhizae to the phytaseactivity of plant roots and utilization of exoge-nously supplied substrate has similarly beeninvestigated in controlled culture and soil-basedexperiments. Phytase activity has been detectedin both arbuscular and ectomycorrhizal fungi(Theodorou, 1971; Bartlett and Lewis, 1973), and


Fig. 15.2. Growth of Nicotiana tabacum in an Alfisol and Spodosol fertilized with different rates ofphosphorus (mg P/kg soil) as either inorganic phosphate or calcium-phytate, or grown without addedphosphorus (No P). (From A. Richardson, 2006, unpublished data.) Shown is the growth of the plants inthe Alfisol at 35 days and shoot dry weight in both soils after 48 days. The Alfisol is a high phosphorus-fixing soil and the Spodosol is a low phosphorus-fixing sand (note the difference in applied rates ofphosphorus). (Details of the soils are reported in George et al., 2005c.) Values are the mean of fivereplicates and the bars show one standard error.

300 Phosphate

Calcium-phytate

200

100Sho

ot d

ry w

eigh

t(m

g/pl

ant)

Sho

ot d

ry w

eigh

t(m

g/pl

ant)

0

No P

Phosphate

Calcium-phytate

200

100

0

300

125

Phosphorus applied (mg P/kg soil)

250 500

No P 15.5 62.5

Phosphorus applied (mg P/kg soil)

125

Calcium-phytate

phosphate

500250

Phosphorus application (mg P/kg soil)

125No P

Nicotianatabacum

Alfisol

Alfisol

Spodosol


in pure culture phytate has been shown to beavailable to a range of mycorrhizal fungi (Antibuset al., 1992; Chen et al., 1999; Sawyer et al., 2003;Midgley et al., 2004). However, McElhinney andMitchell (1993) showed that phytate was hydrol-ysed less effectively than inorganic pyrophosphateor β-glycerophosphate by ectomycorrhizal fungi.Utilization of phytate has also been observed forarbuscular mycorrhizae grown in monoxenic cul-ture with roots (Koide and Kabir, 2000). However,like plant roots, it is evident that phytase frommycorrhizae does not appear to be a significantcomponent of the extracellular activity and gen-erally represents a small component only of thetotal phosphatase activity in mycelia of botharbuscular and ectomycorrhizal fungi (Mousainet al., 1988; Antibus et al., 1992; McElhinney andMitchell, 1993; Firsching and Claassen, 1996;Colpaert et al., 1997).

In the study by Colpaert et al. (1997) phytaseactivity was greater at the surface of ectomycor-rhizal P. sylvestris roots than uncolonized roots.These activities were, however, less than total acidphosphatase and a similar trend was observed forfield-collected roots from a number of differentplant species (Antibus et al., 1997). Whilst the sig-nificance of these observations for the potential ofmycorrhizae to directly utilize inositol phosphatesin soil remains unclear, it is evident that theircontribution to total phytase activity in roots andin soil is likely to be small, as has been reportedfor acid phosphatases (Joner and Johansen, 2000;reviewed by Joner et al., 2000). Despite this, theirrole may be important given that they provide animportant interface between plant roots and thesoil environment.

A number of studies have shown that myc-orrhizae can enhance the uptake of phosphorusby plants when supplied with various sources oforganic matter and added organic phosphorussubstrates (Jayachandran et al., 1992; Joner andJakobsen, 1994, 1995; Perez-Moreno and Read,2000; Tibbett and Sanders, 2002; Baxter andDighton, 2005), although this does not necessarilyimply their direct involvement in mineralization.For example, big bluestem grass (Andropogongeradii) plants colonized with arbuscular mycor-rhizae used phytate (and a range of other organicphosphorus substrates) more effectively than non-mycorrhizal plants (Jayachandran et al., 1992).However, whether the phytate was hydrolysed bythe mycorrhizae or by other soil microorganisms,

and the mycorrhizae simply assisted in its subse-quent uptake, was not determined. In an attemptto separate these aspects, Tarafdar and Marschner(1995) used sterilized soil amended with phytateand inoculated with combinations of both arbus-cular mycorrhizal fungi (Glomus mosseae) and aphytase secreting A. fumigatus. In this study, phos-phorus nutrition of plants was greatest in soilinoculated with both microorganisms and thiswas accompanied with a reduction in soil organicphosphorus. Compartmentalized pots that onlyallow fungal access to soil amended with phytatehave also been used to indicate a mycorrhizal-induced reduction in organic phosphorus in soilimmediately adjacent to the root compartment,suggesting direct utilization of substrate by thefungus (Tarafdar and Marschner, 1994a,b; Fenget al., 2003). On the contrary, using a non-phosphorus-retentive perlite medium, Colpaertet al. (1997) showed that utilization of solublephytate by P. silvestris was poor and that infectionwith two different strains of ectomycorrhizae pro-vided no additional benefit. This occurred despitethe provision of substrate at a relatively high con-centration and observations that phytase activitywas greater on the surface of mycorrhizal roots.

Plant Utilization of EndogenousInositol Phosphates in Soil

Few studies have specifically investigated thebiological utilization of inositol phosphates thatare endogenous to soil, and our understanding oftheir contribution to the phosphorus cycle andrate of turnover in soil is poor. To a large extentthis is due to the lack of access to appropriateanalytical technologies for their direct study insoil and because reactivity of inositol phosphateswith soil constituents is a major factor that restrictstheir availability (e.g. Martin, 1973; Adams andPate, 1992). Despite this, there is emerging evi-dence to suggest that inositol phosphates are bio-logically available, albeit to a limited extent.Early studies showed that inositol phosphateswere mineralized in cultivated soils presumablyas a result of the mixing and subsequent exposureof organic matter to soil microorganisms (Williamsand Anderson, 1968). Subsequently, various micro-organisms in soil that have potential to utilizephytate have been identified (Greaves and Webley,


1969; Cosgrove et al., 1970; Yoon et al., 1996;Richardson and Hadobas, 1997; Idriss et al.,2002; Unno et al., 2005; see Hill and Richardson,Chapter 5, this volume), with many of thesebeing isolated from around plant roots. A num-ber of recent studies have also demonstrated thepresence of phytase-hydrolysable organic phos-phorus (as determined using a range of enzymepreparations that may or may not be specific forhydrolysis of myo-inositol hexakisphosphate; seeRichardson et al., 2005) in various soil fractions,including trace amounts in soil solution andwater extracts, and larger pools within aqueoussoil suspensions (Pant et al., 1994; Shand andSmith, 1997; Hayes et al., 2000a; Hens andMerckx, 2001; Turner et al., 2002a; Toor et al.,2003; Fig. 15.3). Whilst these studies do not pro-vide direct evidence for microbial or plant utiliza-tion of inositol phosphates in soil, they do indicateits potential biological availability.

A more direct approach to the utilization ofinositol phosphates by plants has been the use ofsolution 31P nuclear magnetic resonance (NMR)spectroscopy to investigate the dynamics oforganic phosphorus around roots. Using thisapproach, Chen et al. (2004) and George et al.(2006a) have shown that phosphate monoesterswere depleted from a range of soils and that this

depletion was accompanied by greater acid phos-phatase activity in the rhizosphere. In the study by Chen et al. (2004), depletion was most evidentin soil re-planted with radiata pine (P. radiata) compared to ryegrass, and mineralization of bothmyo- and scyllo-inositol hexakisphosphates wasdemonstrated, with the decline of myo-inositolhexakisphosphate accounting for between 18%and 100% of the observed depletion of the phos-phate monoesters around pine roots (Chen et al.,2004; Turner et al., 2005b). Comparable decreaseswere not observed in soil under the grass.

The greater depletion of inositol phosphatesin soils from radiata pine was considered to bedue to the colonization of the pine roots by ecto-mycorrhizal fungi, as also suggested by others(Liu et al., 2004; Scott and Condron, 2004; Liuet al., 2005). In the study by Scott and Condron(2004) the ectomycorrhizal fungus had access toroot-free compartments of soil where the myceliumdecreased the total extractable soil organic phos-phorus to a similar extent as did the fungus incombination with plant roots. This suggests thatectomycorrhizal fungi may play a dominant role,although the contribution of other soil microor-ganisms to the mineralization of soil organicphosphorus cannot be discounted. Decreases inorganic phosphorus were also smaller in parallel

Fig. 15.3. Phytase-hydrolysable organic phosphorus in soil suspensions of an Alfisol and Spodosol.(From T. George and A. Richardson, 2005, unpublished data.) Phytase-hydrolysable phosphorus wasdetermined using a non-specific phytase on aqueous suspensions (1:10 w/v) of bulk soils either initiallyor after 28 days incubation in a glasshouse either without plants (No plant) or on soil collected from therhizosphere (0–2 mm) of Trifolium subterranean plants that were wild-type transgenic control, orreleased Aspergillus niger phytase (ex::phyA) as an extracellular enzyme. (From George et al., 2005b.)Shown is the concentration of organic phosphorus that was deemed phytase-hydrolysable by incubationof soil samples (5 g) for 24 h with an excess of the non-specific phytase (Sigma Chemical Company,St Louis, Missouri, USA). Bars show one standard error of the mean (n = 4).

0

Phy

tase

-hyd

roly

sabl

e ph

osph

orus

in s

oil s

uspe

nsio

n(µ

g P

/g s

oil)

Alfisol Spodosol

No plantLSD = 2.3(P < 0.05)

After plant growthAfter plant growth Initialsoil

Wild-type

Transgeniccontrol

ex::phyA

4

8

12

16

20Initialsoil

treatments with lucerne (M. sativa L.) and rye-grass that were colonized by arbuscular mycor-rhizal fungi (Scott and Condron, 2004). This isconsistent with other observations that arbuscularmycorrhizae do not appear to play a significantrole in the direct utilization of soil organic phos-phorus above that of the capacity of plant rootsalone (Joner and Jakobsen, 1995; Joner et al., 1995).

Growth and phosphorus nutritionof transgenic plants that release

extracellular phytase

Utilization of inositol phosphates in soil has alsobeen investigated using transgenic plants thatrelease A. niger phytase as an extracellular enzymefrom roots. These plants are useful as they allowcomparisons to be made with control plants that,apart from the release of the phytase, are genet-ically identical. Using phyA-expressing subter-ranean clover, George et al. (2006a) showed thatphosphate monoesters were depleted in the rhi-zosphere of transgenic plants that expressedPhyA and that this depletion accounted for 15%of the total monoester phosphorus (of which~25% could be identified as inositol phosphates)present in an Oxisol. Depletion of phosphatemonoesters phosphates in this soil was accompa-nied by an increase in the alkali-extractable inor-ganic phosphate. When evaluated for growth in arange of different soils, however, the transgenicphyA-expressing subterranean clover did not showimproved phosphorus nutrition over control lines,except when grown in a Vertisol that was high intotal organic phosphorus and water-extractableorganic phosphorus amenable to hydrolysis by asubstrate-specific phytase (George et al., 2005b).In this Vertisol, phosphorus uptake by the trans-genic plants was increased by between 21% and31% over the controls and occurred irrespectiveof whether the soil was pasteurized to minimizethe influence of soil microorganisms.

Interestingly, the lack of phosphorus nutritionresponse of the transgenic plants in other soils(including the Oxisol) occurred despite the pres-ence of significant amounts of inositol phosphatesin each soil, as determined by solution 31P NMRspectroscopy and pools of organic phosphorus thatwere amenable to hydrolysis by phytase (Georgeet al., 2005b). For example, in two of the soils (a low

phosphorus-fixing Spodosol and a high phospho-rus-fixing Alfisol; Fig. 15.2) a significant depletion ofphytase-hydrolysable phosphorus from aqueous soilsuspensions occurred over 28 days in both trans-genic and control plants and in soil that was incu-bated under the same conditions but without plants(Fig. 15.3; T. George and A. Richardson, 2005,unpublished data). Differences were evident in thenet depletion of phytase-hydrolysable phosphorusbetween the two soils and greater depletionoccurred in the rhizosphere of the transgenic linewhen grown in the Spodosol. These results indicatethat either the measure of ‘phytase-hydrolysable’phosphorus in these soils was not a good indicatorof the availability of inositol phosphates to plants,or that microbially-mediated mineralization was adominant process in these soils and that any benefitfrom plant-produced phytases was consequentlyminimal. This is consistent with the lack of growthresponse of transgenic subterranean clover andtobacco plants when grown in these two soils(George et al., 2004, 2005c) and highlights the needto better understand the importance of substrateavailability and its interaction with microorganismsand plant roots in different soils.

Irrespective of this, plant-exuded phytase canbe significant for the phosphorus nutrition ofplants in these soils, because growth of transgenicplants was enhanced over that of control plantswhen the soils were fertilized with either phytateor inorganic phosphate (George et al., 2004,2005c). The response of plants to fertilization withinorganic phosphate (14–32% and 20–50%increase in shoot phosphorus content over wild-type controls for tobacco and subterranean clover,respectively) is of particular interest and suggeststhat phosphate addition may increase the avail-ability of inositol phosphates to plants. This mightoccur through either displacement of adsorbedinositol phosphates, given that the counter-reac-tion (i.e. displacement of phosphate by inositolphosphates) has been observed (Anderson et al.,1974; Helal and Dressler, 1989), or through denovo synthesis by soil microorganisms.

Using a 33P tracer, George et al. (2006a)demonstrated the rapid incorporation of labelledphosphate into alkali-extractable organic phos-phorus, including microbial phosphorus and apool that was amendable to hydrolysis by anon-specific phytase. Based on changes in specificactivity of these pools it was further evident thattransgenic plants that expressed phyA (compared


to control plants) preferentially depleted phos-phorus from recently synthesized organic phos-phorus. Collectively, these results suggest thatmicrobial synthesis of inositol phosphatesoccurred in this soil and that plants that releasedthe microbial-derived phytase had greater accessto this source of phosphorus. However, morework to confirm this hypothesis is required andthere is a need to further investigate the dynam-ics of inositol phosphates in soil, particularly inrelation to the activity of soil microorganisms.Considerable variation in the biochemical prop-erties of microbial phytases has been established(Wyss et al., 1999; Lassen et al., 2001; see Greiner,Chapter 6, this volume) and differential inter-action of these various phytases in soil and theireffectiveness in hydrolysing inositol phosphateshas been demonstrated (George et al., 2005a,2006b; see George et al., Chapter 14, this volume).

Conclusions and Future ResearchDirections

Despite the abundance of inositol phosphates in soil,our understanding of their biological availability andcontribution to the soil phosphorus cycle remainsincomplete. It is evident that inositol phosphates areless available than phosphate diesters and otherphosphate monoesters and are considerably lessavailable than inorganic phosphate. Whilst this maylargely be a consequence of the reactivity of inositolphosphates with soil constituents, it is also apparentthat many plants do not have an innate capacity todirectly utilize inositol phosphates in soil and appearto be dependent on microorganisms for their hydrol-ysis. This raises a number of interesting questionsconcerning the biological relevance of inositolphophate–phytase interactions in soil. In particular,it is paradoxical that plants have not evolved an abil-ity to release phytase and directly utilize inositolphosphates, given the abundance of inositol phos-phates in soils. Plants have evolved a wide range ofother mechanisms to acquire soil phosphate whengrown under conditions of low phosphorus availabil-ity. Either the lack of available substrate in soil solu-tion has precluded selection pressure for such a traitto evolve in plants, or plants have evolved to rely onmicroorganisms for the hydrolysis of inositol phos-phates in soil environments and within the rhizos-phere (e.g. Unno et al., 2005). Obviously there needs

to be further investigation into such possibilitiesusing a much wider range of plant species from dif-ferent ecosystems, in addition to the few agriculturalspecies that have been examined to date. Futureresearch also needs to specifically address the follow-ing issues:

● A better understanding of how plants might utilizeinositol phosphates in soil. This will require moredetailed assessment of the role of phytasesproduced by mycorrhizae and other soilmicroorganisms and how they interact withplant roots in the rhizosphere. These studieswill need to consider the interaction of the dif-ferent phytases with the various stereoisomersof inositol hexakisphosphate that are found insoil (see Turner, Chapter 12, this volume).This should involve compartmentalizedexperimental systems that allow controlledaccess by either roots or fungal hyphae to soil,combined with a wider range of differentplant species, including transgenic plants thathave novel ability to release different phytasesfrom their roots, along with manipulation ofsoil microbial populations. Such experimentsalso need to more closely resemble field situa-tions (e.g. Schweiger and Jakobsen, 2000; Liuet al., 2005) and use analytical procedures suchas solution 31P NMR spectroscopy that allowinositol phosphates to be appropriately identi-fied and quantified separately from other con-stituents of the soil organic phosphorus.

● The development and application of analytical proce-dures that measure ‘biologically relevant’ pools ofinositol phosphates in soil. At present we have lit-tle understanding of which components ofinositol phosphates in soil are amenable tohydrolysis by phytases. Inositol phosphatesdissolved in soil solution would be expectedto be most available to plants and microor-ganisms, yet we have little information tosupport this. Whilst techniques such as NMRspectroscopy provide valuable insight intothe total inositol phosphate content of soil,they currently provide little information ontheir biological availability. There is the needtherefore to develop extraction or fractiona-tion procedures that allow ‘meaningful’ poolsof organic phosphorus to be identified(Turner et al., 2005a). Likewise, a betterunderstanding of the biological relevance ofphytase-hydrolysable pools, as determined by



enzyme hydrolysis assays, is required. Todate these too have proven to be of limitedvalue in relation to the apparent availabilityof inositol phosphates to plants or microor-ganisms. Although transgenic plants thatexpress phytase have shown significant phos-phorus nutrition response in soils that haveboth large concentrations of total and water-extractable organic phosphorus that isamenable to hydrolysis by phytase (Georgeet al., 2005b), these soil properties alone werenot reliable indicators of the potential forplant response. Moreover, the understandingof how various chemical and physical attrib-utes of soil interact with the availability ofinositol phosphates is poor.

● Assessment of the fate of inositol phosphates in soiland factors that contribute to their synthesis anddegradation within different components of soil bio-logical systems. This will require quantitativeanalysis of inositol phosphate turnover insoil (e.g. using radioactive substrates) andcapacity to differentiate ‘newly’ synthesizedcompounds (e.g. by microorganisms in therhizosphere) from more stable forms thatexhibit greater resistance to mineralization.

Recent observations showing differentialinteraction of various phytases with a rangeof metal ion–associated soluble and precipi-tated forms of phytate are significant (Tanget al., 2006), but such studies need to beextended to soil environments. The abilityof microorganisms and plant roots to accessinositol phosphates from these more recalci-trant forms, and to modify the chemicalenvironment for its hydrolysis through therelease of various exudates (e.g. organicacids; Hayes et al., 2000a; Tang et al., 2006)is important. Similarly, the fate of inositolphosphates that enter soil–plant systemsthrough the application of animal manuresand other organic phosphorus residuesneeds to be addressed.

The contribution that inositol phosphates in soilmake to the phosphorus nutrition of plants there-fore remains somewhat uncertain and there is stillmuch to learn concerning the biological interac-tions of inositol phosphates in terrestrial environ-ments. This is important given the predominanceof inositol phosphates in both agricultural and natu-ral ecosystems.

References

Adams, M.A. and Pate, J.S. (1992) Availability of organic and inorganic forms of phosphorus to lupins (Lupinusspp.). Plant and Soil 145, 107–113.

Alvarez, M., Godoy, R., Heyser, W. and Härtel, S. (2005) Anatomical–physiological determination of surfacebound phosphatase activity in ectomycorrhizae of Nothofagus obliqua. Soil Biology and Biochemistry 37, 125–132.

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Xiao, K., Harrison, M. J. and Wang, Z.Y. (2005) Transgenic expression of a novel M. truncatula phytase gene resultsin improved acquisition of organic phosphorus by Arabidopsis. Planta 222, 27–36.


Yoon, S.J., Choi, Y.J., Min, H.K., Cho, K.K., Kim, J.W., Lee, S.C. and Jung, Y.H. (1996) Isolation and identifi-cation of phytase-producing bacterium, Enterobactor sp. 4, and enzymatic properties of phytase enzyme. Enzymeand Microbial Technology 18, 449–454.

Zhang, C. and McManus, M.T. (2000) Identification and characterization of two distinct acid phosphatases in cellwalls of roots of white clover. Plant Physiology and Biochemistry 38, 259–270.

Zimmermann, P., Zardi, G., Lehmann, M., Zeder, C., Amrhein, N., Frossard, E. and Bucher, M. (2003)Engineering the root–soil interface via targeted expression of a synthetic phytase gene in trichoblasts. PlantBiotechnology Journal 1, 353–360.


16 Inositol Phosphates in AquaticSystems

Ian D. McKelvieWater Studies Centre and Chemistry Department, School of Chemistry, Monash

University, Clayton, Victoria 3800, Australia

Although there is a considerable body of researchon the abundance and behaviour of inositolphosphates in soils, much less is known regardingtheir behaviour in aquatic systems. This stemspartly from the emphasis of aquatic research onthe detection and measurement of molybdatereactive species in waters as a surrogate measureof bioavailable phosphorus. Inositol phosphates,like many other organic phosphate species, donot react with molybdate; consequently, theyhave been largely relegated to the fraction ofphosphorus that is considered bio-unavailable,refractory and immobile. This view is overly sim-plistic and this chapter considers the knownsources of inositol phosphates and likely transportpaths and transformations in aquatic systems(Fig. 16.1; Turner et al., 2002b).

The dominant inositol phosphate in soilsand sediments appears to be myo-inositol hexak-isphosphate, with the lower-order inositol phos-phates occurring only as intermediates in eitherhydrolytic or biosynthetic sequences. Thischapter will therefore focus on the behaviour ofmyo-inositol hexakisphosphate in aquatic sys-tems. Suggested mechanisms for the release andtransport of both inorganic and organic phos-phorus from sediments are reviewed, and somespeculative interpretation of the release, hydroly-sis and bioavailability of inositol phosphates isoffered.

Sources of Inositol Phosphates inthe Aquatic Environment

Inositol phosphates in aquatic systems are thoughtto originate from external, terrestrial sources suchas soil particles and plant matter, or from internalsources such as algae and macrophytes. In gen-eral, plants contain only myo-inositol hexakisphos-phate; so it seems likely that the scyllo-, D-chiro-and neo-inositol phosphates are of microbial origin(Cosgrove, 1980), formed by epimerization fromeither myo-inositol or its hexakisphosphate(L’Annunziata, 1975). The phosphorylated inosi-tol stereoisomers are discussed in detail elsewherein this volume (see L’Annunziata, Chapter 4, andTurner, Chapter 12).

Weimer and Armstrong (1979) studied thecomposition of inositol phosphates in severalspecies of aquatic plants, algae and sediments oftwo fresh water lakes in Wisconsin, USA. Foraquatic macrophytes and angiosperms, theyfound that lower-order esters of myo-inositolphosphate (i.e. tetrakisphosphate to monophos-phate) were present in greater amounts than thehexa- and pentakisphosphate esters. They alsonoted that the ratio of higher-order to lower-order inositol phosphates for catchment soils wasgreater than that found in the sediments, andsuggested that this was due either to hydrolysisof soil inositol phosphates during transport into

262 I.D. McKelvie

the lake or to a much larger contribution of inos-itol phosphates from plant sources than expected.They concluded, however, that inositol phos-phates in these lakes were derived approximatelyequally from algal primary production and catch-ment inputs, rather than from aquatic plants.

These early findings were subsequently ques-tioned, because the extraction conditions used (pH<1, 110°C, 6 h) may have dephosphorylatedhigher inositol phosphates (Potman and Lijklema,1983). However, there are other reports of in vivobiosynthesis of inositol phosphates by aquaticplants, so this may be an important autochthonoussource. For example, the aquatic angiospermWolffiella floridana was found to convert a high pro-portion of radiolabelled myo-inositol to inositolhexakisphosphate and lower inositol phosphatesrather than to cell wall polysaccharides (Robertsand Loewus, 1968). Significant amounts of myo-inositol hexakisphosphate were also found in theduckweed Spirodela polyrhiza during a developmen-tal stage (Brearley and Hanke, 1996a), and thesequence of phosphorylation in this process wassubsequently described (Brearley and Hanke,1996b). Phosphorylation of myo-inositol to producemyo-inositol hexakisphosphate has also beenreported to occur in the lesser duckweed Lemnaminor (Inhulsen and Niemeyer, 1978) and the slimemould Dictostelium (Stephens and Irvine, 1990).

Inositol phospholipids and lower-orderinositol phosphates have also been implicated inphytoplankton metabolism and structure (Oku

and Kamatani, 1995). However, in a study of thecoastal sediments in Tokyo Bay, Japan, Suzumuraand Kamatani (1995b) found very little inositolphosphate in zooplankton and algae and reportedthat the major source of inositol phosphates wassoils from surrounding catchments. This conclu-sion was based on the observation that the orderof abundance of the myo-, scyllo-, and chiro-inositolphosphates were the same in sediments as in ter-restrial and river suspended particulate samples.Hence, if biosynthesis were a major source ofinositol phosphates in these sediments, it wouldbe likely that varying ratios of inositol phosphatestereoisomers would be detected.

The input of inositol phosphates from themanures of monogastric animals (poultry, swine)into aquatic systems is also a potentially largesource of inositol phosphate, especially fromcatchments containing intensive production (e.g.feedlot farming). The manures of monogastricagricultural animals can contain high concentra-tions of myo-inositol hexakisphosphate becausesuch animals have low levels of intestinal phytase(e.g. Maguire et al., 2004), and for this reasonphytase is increasingly used as a feed additive toimprove utilization of inositol phosphates and toreduce the requirement for inorganic phosphatesupplements in animal diets (Valaja et al., 1998;Bedford, 2000; Juanpere et al., 2004; see Lei andPorres, Chapter 9, this volume). Improvement indietary phosphorus availability through the useof phytase also appears to be beneficial by reduc-

Fig. 16.1. Suggested sources, pathways and transformations of inositol hexakisphosphate (IP6) inaquatic ecosystems. (Modified from Turner et al., 2002b.)

Inositol Phosphates in Aquatic Systems 263

ing the amount of inorganic phosphorus excretedand added to catchments from manures (Pennet al., 2004; see Leytem and Maguire, Chapter10, this volume). An extension of this approachhas been the development of transgenic pigs thatproduce phytase in their saliva (Golovan et al.,2001). This enables them to digest phytate in thediet and is claimed to reduce faecal phosphorusoutput by up to 75%.

Manure from poultry and swine not fed phy-tase-amended diets is likely to contain high concen-trations of myo-inositol hexakisphosphate (althoughsee Leytem et al., 2004), but it is poorly soluble andwould likely be strongly bound to soils (Turner andLeytem, 2004). However, despite its low solubility,transport into streams in either colloidal or particu-late forms might reasonably be expected underoverland flow conditions (Turner, 2005).

Physicochemistry of InositolPhosphates in Aquatic Systems

myo-Inositol hexakisphosphate is resistant to chemi-cal hydrolysis, especially under alkaline conditions.The maximum rate of hydrolysis occurs near pH 4(Cosgrove, 1980) and decreases to a minimum atpH 0–1. Less than 50% hydrolysis was achieved atpH < 0 and 100°C for 6 h, (Cosgrove, 1980),whereas at 50°C hydrolysis was <20% after 35 days(Potman and Lijklema, 1983). This behaviour hasimportant implications for both the extraction ofinositol phosphates from sediments and soils, andthe analysis of the organic phosphorus componentof sediments, pore waters and the overlying watercolumn. Extraction with strong acids and highertemperatures for extended periods introduces therisk of myo-inositol hexakisphosphate hydrolysis(Potman and Lijklema, 1983, Turner et al., 2005a),which has been cited as a possible explanation forthe high concentrations of lower-order inositolphosphates reported in lake sediments by Weimerand Armstrong (1979). On the other hand, diges-tions for total and total filterable phosphorus deter-mination using nitric acid alone or nitric andsulphuric acids may give incomplete recovery ofmyo-inositol hexakisphosphate, especially if typicaldigestion times of less than 2 h are used. For com-plete conversion, inclusion of an oxidizing agentsuch as peroxydisulphate or hydrogen peroxide inthe digestion reagent is recommended (Benson et al.,

1996). This also highlights the need for adequatevalidation of the extraction, digestion and detectionstages in the determination of inositol phosphatesusing appropriate model organic phosphorus com-pounds (Kérouel and Aminot, 1996).

Abiotic reactions of inositol phosphates withcharged surfaces and polyvalent cations are dis-cussed in detail elsewhere in this volume (see Celiand Barberis, Chapter 13) and only a briefoverview of aspects relevant to aquatic systems isprovided here. myo-Inositol hexakisphosphate read-ily complexes multivalent metal ions, and the orderof relative complex stabilities is thought to be:Cu2+> Zn2+ > Ni2+ > Co2+ > Mn2+ > Fe3+ > Ca2+

(Cosgrove, 1980). It also forms stable surface com-plexes with minerals such as goethite and Fe(OOH)(De Groot and Golterman, 1993), and with clayminerals such as illite and kaolinite. Adsorptionmay also disperse clay minerals by alteration ofsurface charge (Celi et al., 1999). The sorption ofmyo-inositol hexakisphosphate on goethite wasobserved to depend both on the nature of dissolvedcations present and the pH. In the presence ofpotassium ions, adsorption was pronounced at lowpH, but as pH increased the adsorption decreasedin response to the increasing charge density of myo-inositol hexakisphosphate (−8 at pH 5.5).Adsorption on to goethite (Fe(OOH)) also displacedphosphate that was already adsorbed, and pre-vented further adsorption of phosphate. As wasthe case for the clay minerals, myo-inositol hexak-isphosphate adsorption had a pronounced effect onthe surface charge, resulting in dispersion ofgoethite particles. When calcium ions are present,however, adsorption continues to occur even athigher pH, apparently due to precipitation of cal-cium-phytate complexes. In this case, the effect ofmyo-inositol hexakisphosphate adsorption is insuffi-cient to cause particle dispersion, and aggregationof goethite occurs in the presence of Ca2+ (Celiet al., 2001; see Celi and Barberis, Chapter 13, thisvolume).

De Groot and Golterman (1993) also studiedthe effect of iron(III) reduction on myo-inositolhexakisphosphate adsorbed on goethite andreported that rather than being released in a solu-bilized form, myo-inositol hexakisphosphateremained bound as insoluble Fe4-phytate. Calcite isalso reported to have a high capacity for retentionof myo-inositol hexakisphosphate; this appears toinvolve a combination of adsorption and the com-plexation of calcium ions, accompanied by the dis-

264 I.D. McKelvie

solution of calcite and the precipitation of Ca3-phy-tate (Celi et al., 2000).

These adsorption and aggregation/disper-sion effects have important implications for theaccumulation of inositol phosphates in soils and colloidal transport in waters. The observed phy-sicochemical behaviour of myo-inositol hexakis-phosphate suggests that in marine and freshwaters it should be stable, complexed and/orprecipitated by major ions such as calcium, mag-nesium, iron and manganese, or bound to thesurfaces of suspended particles and thus immobi-lized in the sediments (De Groot and Golterman,1993). In addition, inositol hexakisphosphate isthought to be associated with high-molecularmass humic acids (Hong and Yamane, 1980;Golterman et al., 1998).

Occurrence of Inositol Phosphates inAquatic Systems

Waters

A commonly held view is that inositolhexakisphosphate exists in aquatic systems asan insoluble, refractory, immobile and biologicallyunavailable phosphorus species. Inositol hexak-isphosphate is not hydrolysed by exocellular alkalinephosphatase or phosphodiesterase (Turner et al.,2002a), and is therefore regarded as biologicallyunavailable to phytoplankton. Thus it might beexpected that the concentration of inositol phos-phate in solution in both pore and overlying waterswould be very low, or even undetectable. However,Eisenreich and Armstrong (1977) used a combina-tion of alkaline bromination and gel filtration toperform selective detection of inositol phosphates,and reported that there was between 3 and 15 µgP/l of this organic phosphorus species in the watersof Lake Mendota, Wisconsin, USA, which repre-sented between 20% and 30% of the total filterablephosphorus present (Table 16.1). Similarly, substan-tial concentrations of high-molecular mass phos-phorus, with similar elution times as inositolhexakisphosphate on a gel filtration column, havebeen detected in the filterable fraction of waters andsediment pore waters (Fig. 16.2) from south-eastAustralia (McKelvie et al., 1993; McKelvie, 2005).Concentrations of this fraction in a range of waterswere between undetectable and 52 µg P/l, and in

some cases comprised most of the filterable phos-phorus present. Although the resolution of the gelfiltration separation was limited, the dominant high-molecular mass phosphorus peak always coincidedwith that for an authentic standard of myo-inositolhexakisphosphate; so it is reasonable to assume thatthis peak consisted predominantly of higher-orderinositol phosphates.

Others have used hydrolytic techniquesbased on the enzyme phytase to hydrolyse organicphosphorus in waters and sediment extracts(Cooper et al., 1991). For example, it was shownthat up to 50% of the organic phosphorus in thewaters of two small mesotrophic and hypereu-trophic lakes was amenable to hydrolysis by phy-tase, and on the basis of gel filtration separations,that this organic phosphorus consisted either ofsoluble inositol phosphates or inositol phosphatesassociated with higher-molecular mass proteins,lipids or fulvic acid (Herbes et al., 1975). The sameenzyme, 3-phytase, was used in an immobilizedform in an automated flow injection system todetermine the concentration of phytase-hydro-lysable phosphorus in waters (McKelvie et al.,1995). Concentrations in a range of waters insouth-east Australia were between 1 and 75 µgP/l, with the higher values being associated withestuarine waters.

The determination of inositol phosphates bythese approaches is, however, more inferentialthan definitive. The specificity of commercial3-phytase preparations is poor; it catalyses hydroly-sis of a wide range of phosphomonoesters andeven some diesters (McKelvie et al., 1995).Consequently, concentrations reported as myo-inositol hexakisphosphate by this approach willoverestimate the true concentration. Similarly, thepoor selectivity of low-pressure gel filtration sepa-rations means that, at best, separated peaks, suchas those shown in Fig. 16.2, represent a size ormass range rather than a single species such asmyo-inositol hexakisphosphate. The use of analyti-cal techniques with high selectivity for inositolphosphates, such as that described by Clarkin et al.(1992) or Suzumura and Kamatani (1993), ispreferable. For example, Espinosa et al. (1999),using high-performance ion exchange chromatog-raphy, obtained highly resolved separations andshowed that myo-inositol hexakisphosphate consti-tuted nearly one-third of the identifiable organicphosphorus compounds present in leachate froma temperate grassland soil.

Inositol Phosphates in A

quatic System

s265

Table 16.1. Indicative concentrations of inositol hexakisphosphate in water, using a variety of estimation methods.

Inositol hexakisphosphate

Fraction of total Sample origin and type (µg P/l) filterable P (%) Method of detection Reference

Frain’s Lake, USA (highly 3.5–12.4 12–47a Phytase hydrolysis, photometric detection of Herbes et al. (1975)eutrophic) reactive phosphate

Third Sister Lake, USA 4.9–10.0(eutrophic)

Lake Mendota, Wisconsin, 3–15 20–30 Alkaline bromination, followed by gel filtration Eisenreich and USA on Sephadex G25 Armstrong (1977)

River (4) and estuarine (5) 1–75 (mean 25) 15 Phytase hydrolysis, flow injection detection of McKelvie et al. (1995)waters, rural Victoria, reactive phosphatesouth-east Australia

Urban river (2) and lake 21–52 (mean 36) 83 Gel filtration on Sephadex G25, flow injection McKelvie et al. (1993)waters (2), Melbourne, detection of reactive phosphate after Australia online photooxidation

Yarra River sediment, Gel filtration on Sephadex G25, flow injection McKelvie (2005)b

south-east Australia detection of reactive P after online photooxidation

Overlying water 27 66Pore water 29–281 (mean 85) 85

aProportion of the total organic phosphorus.bSee Fig. 16.2 for further details.

266 I.D. McKelvie

Sediments

Given the apparent refractory and insolublenature of metal-complexed myo-inositol hexak-isphosphate, it might be expected that sedimentswould contain substantial amounts of phosphorusin this form. This is supported by reports that itmay comprise as much as 80% of the totalorganic phosphorus (Weimer and Armstrong,1977). However, the amount detected will be

highly dependent on the respective efficiency andthe selectivity of the extraction and detectionmethods used (Table 16.2).

Historically, soils and sediments wereextracted with dilute acid to remove calcium-bound phosphate, followed by strongly alkalinemedia such as hot 3 M NaOH to recoverorganic phosphorus, with final precipitationof inositol phosphates as barium salts (reviewedin Turner et al., 2002b). Further isolation of

Fig. 16.2. (a) Low-pressure, gel filtration separations of high- and low-molecular mass organicphosphorus (HMMP and LMMP, respectively). myo-Inositol hexakisphosphate and phosphate eluted at~2.8 and 5.5 min, respectively. (b) Concentrations of phosphorus found in pore waters from Yarra River(Fairfield) sediment. (Redrawn from McKelvie, 2005.)

0 2 4 6 8 0 2 4 6 80

20

40

60 16

12

8

4

0

Overlyingwater

0−1 1−2 2−3 3−4 4−5 5−7.5

Pore water depth (cm)

300

250

200

150

100

50

0

Pho

spho

rus

conc

entr

atio

n (µ

g P

/I)

LMMPHMMP

0−1 cm core segment 3−4 cm core segment

Time (min) Time (min)

(a)

(b)

Inositol Phosphates in A

quatic System

s267

Table 16.2. Reported concentrations of myo-inositol hexakisphosphate in sediments.

Inositol hexakisphosphate

Fraction of Sample origin and type mg P/kg dry wt total P (%) Extraction and analytical methodology Reference

Sediments from ten lakes, – 51–80a 0.3 M NaOH, 25ºC, 16 h, and 0.3 M Sommers et al. (1972)Wisconsin, USA NaOH, 100ºC, 8 h; ion exchange

chromatographySediments from Lake Mendota, 49 12.1 3 M NaOH, 100ºC Weimer and Armstrong (1977)

Wisconsin, USA 38 9.3 1 M NaOH, 60ºC, hypobromite oxidation, barium precipitation

51 12.7 3 M NaOH, 100ºC, hypobromite oxidation, barium precipitation; ion exchange chromatography

Sediments from Tokyo Bay, Japan 1.9–6.2 (mean 3.7) 0.39 (mean) Hypobromite oxidation; ion exchange Suzumura and Kamatani (1993)chromatography; gas chromatography

Riverine and estuarine suspended 2.2–20.4 (mean 8.8) 0.75 (mean) Hypobromite oxidation; ion exchange Suzumura and Kamatani (1995b)solids from Tokyo Bay, Japan chromatography; gas chromatography

Sediments from Tokyo Bay, Japan 0.3–3.1 (mean 1.9) 0.22 (mean) Hypobromite oxidation; ion exchange Suzumura and Kamatani (1995b)chromatography; gas chromatography

Sediments from Lake Wellington, 2.5–14 (mean 6.3) 53 (mean) 25 mM Na-tetraborate, pH 9.2, McKelvie et al. (1993)Australia measured as high-molecular mass

phosphorus using gel filtration with flow injection analysis

Marsh and lake sediments from 24–149 (mean 86) 17.6a (mean) 0.5 M HCl, 30 min, 2 M NaOH, 90ºC, De Groot and Golterman (1993)Camargue, France 30 min, H2SO4, pH < 2, phytase

aProportion of the total organic phosphorus.

268 I.D. McKelvie

myo-inositol hexakisphosphate can be achievedby the use of ion exchange chromatography(McKercher and Anderson, 1968), althoughthis approach lacks selectivity because otherorganic phosphorus species may also be pres-ent. To overcome this, Irving and Cosgrove(1981) used bromine to oxidize all soil organicmatter and organic phosphorus other than thehigher-order inositol phosphates. However, ithas been shown subsequently that while thisprocedure gives good recovery for myo-inositolhexakisphosphate (Weimer and Armstrong,1977), it does not oxidize DNA (Nanny andMinear, 1994).

The efficiency of myo-inositol hexakisphos-phate extraction of soils was shown to improve ifhigher temperatures were employed in thesodium hydroxide extraction step and longertime was allowed for the hypobromite reaction(Hong and Yamane, 1980). To avoid possibledegradation of the sample by high temperatureextraction a milder procedure was recommendedfor sediments (De Groot and Golterman, 1993).This involved extraction with complexing reagentssuch as ethylenediaminetetraacetate (EDTA) anddithionite, followed by extraction of acid-solubleand residual organic phosphorus (containinginositol phosphates) with 2 M NaOH (Fig. 16.3).

Pellet 0 (Total P)

Ca-NTA 0.02 M Dithionite

pH = 7.8−8.0

Na-EDTA 0.05 M

pH = ~8.0

Fe(OOH)~PAmorphous Fe(OOH)

CaCO3~PCaCO3

Pellet I (Organic P)

Extractionof

inorganic-P

0.5 M H+ (HCI or H2SO4)

30 min

Pellet II (ROP = residual organic phosphate)

2.0 M NaOH 90°C

30 min

Pellet III (Rest~P)

NaOHextr~P

H2SO4

pH < 2.0

Fulvic acid~PPhytase

Pellet IV(Humic acid~P)

Phytate~P(Inositol phosphate)

ASOP(Acid-soluble organic P)

Fig. 16.3. Scheme for phosphorus fractionation in sediments, including the determination of inositolhexakisphosphate in the residual organic phosphate component. (Reproduced from De Groot andGolterman, 1993 with permission from Springer-Verlag.)


A further consideration in the selection of anextraction scheme is compatibility with the detec-tion method to be used. Cade-Menun andPreston (1996) reported that extraction with 0.25M NaOH and 0.05 M EDTA is suitable for boththe extraction of organic phosphorus and subse-quent characterization by solution 31P nuclearmagnetic resonance (NMR) spectroscopy. Similarextraction conditions have been used in theextraction and determination of inositol phos-phates from soils (Turner and Richardson, 2004)and manures using 31P NMR (Turner andLeytem, 2004).

Given the ubiquity of inositol phosphates insoils and their known physicochemical behav-iour, it is reasonable to suppose that high con-centrations would be found in sediments, andthat once buried, they would remain bound asmetal precipitates. However, a study of the dis-tribution of inositol phosphates in coastalmarine and estuarine sediments from TokyoBay showed that although riverine suspendedparticulate matter and estuarine sedimentscontained appreciable amounts of inositol phos-phates, the concentrations decreased progres-sively towards the mouth of the bay (Suzumuraand Kamatani, 1995b). Further, inositol phos-phate concentrations were high in surface sedi-ments, but were almost completely absent indeeper layers (Fig. 16.4).

Inferential evidence of the breakdown ofinositol phosphate in sediments was also providedby a study that isolated and quantified inositols inmarine sediments (White and Miller, 1976).While the proportions of unphosphorylated myo-,chiro- and scyllo-inositols were similar with depth,the total inositol concentration decreased. Thiswas attributed to the effects of bacterial actionand leaching on inositol phosphates.

Further studies of Tokyo Bay sediments(Suzumura and Kamatani, 1995a) showed thatunder anaerobic conditions inositol hexakisphos-phate was almost completely mineralized within40 days, while under aerobic conditions it took60 days for ~50% decomposition to occur.Decomposition under anaerobic conditions wasascribed to bacterial hydrolysis, and was con-sidered to be more pronounced in the marineenvironments compared with fresh water environ-ments. Depth profiles of myo-inositol hexakisphos-phate in Yarra River pore waters (measured ashigh-molecular mass phosphorus; Fig. 16.2)showed a similar trend to that in Fig. 16.4(McKelvie, 2005), although the presence of meas-urable myo-inositol hexakisphosphate in deeper sed-iments suggested that the rate of decomposition orremoval is not as fast as that in marine systems.

In a recent paper, Turner and Newman(2005) reported a distinct absence of inositolphosphates in wetland soils and benthic floccu-

Inorganic, organic P (µmol/g )

IP6−P (�10−2 µmol/g )

0

5

10

15

20

25

0

5

10

15

20

25

0 5 10 15 20 0 5 10 15 20

Dep

th (

cm)

IP6

IP6

Org

anic

Org

anic

Inor

gani

c

Inor

gani

c

Fig. 16.4. Vertical distribution of inositol hexakisphosphate (IP6), inorganic phosphate and organicphosphorus from two cores in Tokyo Bay, Japan. (Reproduced from Suzumura and Kamatani, 1995bwith permission from Elsevier.)

270 I.D. McKelvie

lent from the Florida Everglades, USA, with themajority of the organic phosphorus being presentas phosphate diesters. A possible explanation forthis observation is that either there is no terres-trial source of myo-inositol hexakisphosphate tothis system or the rates of decomposition andremoval are much greater than the rate of supplyfrom terrestrial sources.

On the basis of these few examples, it is evi-dent that inositol hexakisphosphate is not asrefractory or immobile as previously thought,and that remobilization and mineralization ofthis organic phosphorus species may comprise animportant source of bioavailable inorganic phos-phate in sediment pore waters and the overlyingwater column.

Phosphorus Remobilization at theSediment–water Interface

While sediments are a major sink for both inor-ganic and organic phosphorus, there is stronginterest in the biological, physical and chemicalconditions that favour the release of the internalload of phosphorus from sediments back intothe overlying water. The importance of theinternal cycling and release of bioavailableinorganic phosphorus from sediments as a con-

tributor to algal blooms and eutrophicationhas long been appreciated. However, theemphasis has mostly been on bioavailable inor-ganic phosphate, and the role of organic phos-phorus species in this process has largely beenignored.

Phosphates in sediments may be eithersorbed to, or co-precipitated with, metal hydro-xyoxides, clay minerals and calcium carbonate,or bound to humic substances. In oxygenatedwaters, the sediments are covered with an oxi-dized microzone of iron(III) (e.g. Fe(OOH)) thatwill sorb phosphorus from overlying waters andact as a surface barrier, preventing diffusion ofphosphorus from the sediment pores into theoverlying water (Wetzel, 1999). This sectionsummarizes the proposed mechanisms for sedi-ment phosphorus remobilization. This processhas two general components: (i) the release ofphosphorus species from the particulate phaseinto the pore water, which in the case of organicphosphorus may involve either desorption ormineralization; and (ii) the transport of thisphosphate-enriched pore water into the overly-ing water. The processes involved in both com-ponents is shown schematically in Fig. 16.5(Boström et al., 1982; Wetzel, 1999; Golterman,2001) and this section attempts to reconcilethese with the observed behaviour of inositolphosphates.

Fig. 16.5. Schematic diagram showing important processes involved in the release of phosphorus fromsediments. (Modified from Boström et al., 1982 and Wetzel, 1999.)

TRANSPORTMECHANISMS

PHYSICOCHEMICALMOBILIZATION

Diffusion

Dissolved phosphorus

Particulate phosphorus

Desorption Dissolution Ligandexchange

Bioturbation Gas ebullition

Water

SEDIMENT

MICROBIALMOBILIZATION

Enzymatichydrolysis

Wind-inducedturbulence


Anaerobic conditions

Phosphate is released from sediments underanaerobic conditions, and the most frequentlyadvanced explanation for this behaviour is thesolubilization of bound phosphates when iron(III)is reduced to soluble iron(II) (Einsele, 1936;Mortimer, 1941, 1942). Despite the popularity ofthis direct reduction or oxygen control model(Fig. 16.6), Golterman (2001) has argued thatthere is little evidence to support it. Instead, heand others have proposed that sulphate reductionunder strongly reducing conditions would lead tothe formation of insoluble FeS from Fe(OOH),thus indirectly releasing adsorbed phosphatespecies from Fe(OOH)≈P (Fig. 16.6). Severalstudies provide strong evidence in support of thismechanism for phosphate release (Caraco et al.,1989; Roden and Edmonds, 1997; Rozan et al.,2002). Given that myo-inositol hexakisphosphateis adsorbed to Fe(OOH) in preference to phos-phate (Celi et al., 2001), its release from anoxicsediments by this mechanism is perhaps morefeasible than that of solubilization of Fe(OOH),especially as iron(II)-phytate is reportedly insol-uble (De Groot and Golterman, 1993). Goltermanet al. (1998) also suggested that anaerobic fermen-tation of phytate in sediments is a possible source

of phosphate release, which might explain thedramatic decrease in myo-inositol hexakisphos-phate in anaerobic sediments of Tokyo Bay(Suzumura and Kamatani, 1995b) and theabsence of inositol phosphates in Florida wet-lands (Turner and Newman, 2005).

Aerobic conditions

Phosphate is released from sediments under aero-bic conditions, especially in shallow, non-strati-fied systems that are well oxygenated (Boströmet al., 1982). Bacterial mineralization of organicphosphorus through hydrolysis of phosphateesters by enzymes such as alkaline phosphatase isthought to be an important remobilization mech-anism (Fig. 16.7). These extracellular phosphohy-drolytic enzymes are produced by algae andbacteria and are reported to have high activity inthe suspended particulate and sediment phases(Boon, 1989). However, inositol hexakisphosphateis not amenable to hydrolysis by alkaline phos-phatase (McKelvie et al., 1995) and many algae,while possessing phosphomono- and diesteraseactivity, show no phytase activity (Whitton et al.,1990, 1991). Consequently, inositol phosphate

Fig. 16.6. Possible mechanisms for the release of phosphorus species from anaerobic sediments inresponse to sulphate reduction and direct reduction of iron(III). Diss. P = dissolved phosphorus; SRB =sulphate-reducing bacteria; IRB = iron-reducing bacteria. Fe(OOH)˜P represents iron-associatedphosphorus species (phosphate, organic phosphorus). (Modified from Roden and Edmonds, 1997.)

272 I.D. McKelvie

species have been considered bio-unavailable.However, some cyanobacteria such as Calothrixexhibit phytase activity (Livingstone et al.,1982, 1983), which may represent an importantmechanism for the hydrolysis of myo-inositolhexakisphosphate. Similarly, a number of othermicroorganisms in aquatic environments havethe capacity to utilize phytate (see Hill andRichardson, Chapter 5, this volume).

Organic phosphorus mineralization may alsooccur under aerobic sediments due to bacterialrespiration (Fig. 16.7). Oxygen is used as an elec-tron acceptor in this process, although nitratemay also be utilized as oxygen becomes depleted.Release of inorganic phosphate under net aerobicconditions has been reported, but it is suggestedthat even at moderately oxidizing redox potentialsanaerobic microzones will occur on the sedimentsurface and that either direct or indirect reductioncan occur at these sites (Boström et al., 1982).

In shallow water bodies that are well lit,sediments may be covered by benthic algal films.Photosynthetic production of oxygen by thismicro-phytobenthos will increase the thickness ofthe oxidized microzone and reduce the flux ofphosphorus from pore waters. They may act as aphysical barrier that retards upwards diffusion ofpore water, although they can enhance theuptake of phosphorus from overlying waters intothe sediments (Underwood, 2001). In the dark,

benthic algal films may also release phosphorusfrom sediment due to respiratory breakdown oforganic phosphorus (Graneli and Sundback, 1985).Given the complexity of these microphytobenthosassemblages, it is not improbable that they mightproduce phytase as a means of utilizing myo-inositolhexakisphosphate and other organic phosphorussubstrates, although this has yet to be tested.

pH and ligand exhange processes

As sediments become anoxic their pH decreasesdue to the increase in dissolved carbon dioxide.In eutrophic hardwater systems this can solubilizeapatite and release associated phosphate(Golterman, 1998). On the other hand, increas-ing the pH decreases the sorption capacity ofiron(III) hydroxyoxides, and hence the amount ofphosphate or organic phosphorus adsorbed. Thisbehaviour may be due to ligand competition byhydroxyl groups and two possible mechanismshave been proposed (Fig. 16.8). Sediment releaseof inositol hexakisphosphate by this mechanism isquite feasible given that sodium hydroxide is suc-cessfully used to extract myo-inositol hexakisphos-phate from soils and sediments. Data fromRippey (1977) cited in Boström et al. (1982) showthat higher rates of phosphorus release from

Fig. 16.7. Possible mechanisms for the release of phosphorus species from sediments in aerobic,shallow water systems. The scale of the arrows is not representative of flux magnitude.


sediments occurred from about pH 8.2 upwards.This corresponds to the pH of sea water, andsuggests that ligand exchange associated withincreasing pH in Tokyo Bay might explain someof the loss of myo-inositol hexakisphosphate frommarine sediments there. However, the increas-ingly negative charge on inositol hexakisphos-phate (~ −9 at pH 8.2) will tend to oppose, if notentirely negate, ligand exchange as a possiblerelease mechanism for myo-inositol hexakisphos-phate. Bacteria from lake sediments are alsoreported to produce organic acids in conjunctionwith carbohydrate metabolism and growth inaerobic systems. These organic acids cansequester metal ions in metal–phosphate com-plexes (e.g. FePO4) resulting in the solubilizationof the phosphorus species (Boström et al., 1982).

Salinity

Changes in salinity have been observed to releaseboth inorganic and organic phosphorus from sedi-ments, most probably through a combination ofthe lysis of bacterial cells and ligand exchange(Gardolinski et al., 2004). It was noted that signifi-cant organic and inorganic phosphorus release (10µg P/l) occurred at salinity values >10 on the prac-tical salinity scale, and that this was followed byrapid hydrolysis and release of bioavailable reactivephosphate. The potential for inositol phosphateremobilization by this route is still unknown.

Physical and biological sedimentperturbation

In shallow waters, wind- and tidal-induced turbu-lence will cause suspension of surficial sediment,which favours release of phosphorus-rich porewater back into the water column over the muchslower diffusion process. However, phosphorusmobilized in this manner may be rapidly read-sorbed to suspended particulate matter (Holdrenand Armstrong, 1980). Other physical processescapable of disturbing the sediment include gasebullition (e.g. nitrogen gas as part of denitrifica-tion). The sediment surface layer may also be dis-rupted through bioturbation and bioirrigation byorganisms such as tubificid worms, chironomidsand benthivorous fish.

Future Research

There is little appreciation of the magnitude ofinositol phosphate transport within the aquaticenvironment. Given that inositol phosphates canconstitute a sizeable fraction of the total phos-phorus in soil particles, riverine transport of sus-pended sediments is a potentially importantsource of phosphorus in estuaries and coastalwaters. This is exemplified using data from theUK. Taking 133 mg P/kg as an average concen-tration of myo- and scyllo-inositol hexakisphos-phates for soils (Turner et al., 2003, 2005b) and

Me O P

O

OH

O R + OH− Me O H P

O

OH

O R−O+

(a)

Fe

Fe

O

OP

O

O R+ OH−

Fe

Fe

OH

OH

−O

−OP

O

O R+

(b)

—

— — — — —— — — —

—

— ——

—— —

Fig. 16.8. Possible hydroxyl ligand exchange mechanisms that would account for phosphorus removalat higher pH (a) from Lijklema (1977) and (b) from Andersen (1975) reported in Boström et al. (1982). Rrepresents either a proton or an organic moiety.

274 I.D. McKelvie

an annual suspended solids load for UK rivers of45 × 109 kg (Littlewood and Marsh, 2005), anestimated annual load of 6 million kg of inositolhexakisphosphate-phosphorus can be calculated.This represents a relatively minor amount ofphosphorus compared with the equivalent annualload of dissolved phosphate of ~370 million kg(Littlewood and Marsh, 2005). However, as thedissolved phosphate load includes a considerablecontribution from point sources (e.g. Sharpley andWithers, 1994), such as sewage and industry, theinositol phosphate load probably accounts for asubstantial component of the phosphorus exportfrom diffuse sources.

Transport of inositol phosphates to aquaticecosystems is of potential significance, because itis becoming increasingly evident that they are notrefractory, immobile phosphorus species as wasonce thought. The detection of myo-inositol hexa-kisphosphate in the pore and overlying waterssuggests that there is diffusional transport fromsediments, and that the released compounds maybe converted to more bioavailable forms in thepresence of hydrolytic enzymes. The literatureabounds with reference to the association of inos-itol phosphate with higher-molecular weightorganic matter such as humic material, but as yetthere is no clear understanding of the nature ofthese interactions or their importance in thecycling of inositol phosphates.

To a large extent, the study of inositol phos-phates in aquatic systems has been hampered bythe absence of suitable and accessible techniquesfor their analysis and detection (Turner et al.,2002b). For example, it is unclear whether myo-inositol hexakisphosphate detected in overlying

waters is present in true solution or a colloidalform. This question will probably not beanswered by measurements based on gel filtra-tion or ion exchange after hypobromite oxida-tion, and the use of less invasive preparation andseparation techniques will be required.

Similarly, studying the role of phytase in thehydrolysis of inositol phosphates in sediments andwaters has been complicated by the lack of anartificial substrate that would allow straightfor-ward measurement of phytase activity (Turneret al., 2002b). The recent development of a `teth-ered’ inositol phosphate compound that can beused as a substrate in activity measurementsassists greatly in elucidating the role of bacteriaand algae in hydrolysing myo-inositol hexakispho-sphate (Berry and Berry, 2005).

Further elucidation of the behaviour of myo-inositol hexakisphosphate at the sediment–waterinterface will also require improvement of sam-pling techniques other than that offered by thestraightforward collection of sediment cores andpore waters. Devices such as benthic chambers forflux measurements across the sediment–waterinterface, and diffusive gradients in thin films(DGT) (Zhang et al., 1998) with binding phasesspecifically designed for organic phosphorus,would provide valuable information in this respect.

Acknowledgement

I am indebted to Dr Philippe Monbet for his crit-ical comments and assistance in redrafting somediagrams.

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279

Index

Abiotic reactions of inositol phosphatesin aquatic systems 263–264in soil 207–217see also Adsorption

Acid dissociation constants (pKa) 17–18, 214, 215Acid hydrolysis, soil extracts 41, 42, 49Acid phosphatases 244

see also Purple acid phosphatasesAcid protease 137–138Adenosine diphosphate–glucose pyrophosphorylase

(AGP) 119Adsorption (sorption)

inositol phosphates in aquatic systems 263–264inositol phosphates in soils 207–213

effects on surface properties 216–217effects on utilization by plants 245role of calcium carbonate, clays and

organic matter 211–213role of iron and aluminium oxides

208–211solution characteristics affecting 214–216

phytases in soils 223–225Aerobic conditions, sediments 271–272Agranoff’s turtle 4, 5Algal films, benthic 272Alkaline phosphatase 271Alkaline phytases 102, 103Alum (aluminium sulphate) 163Aluminium ions

inositol phosphate hydrolysis and 175, 177–178inositol phosphate reactivity 172–173

Aluminium oxides, soil adsorption and 208–211Amoebae 195Anaerobic conditions

in aquatic systems 269, 271inositol phosphate hydrolysis 192, 269–270

phytase synthesis 83, 84, 85phytate-degrading microorganisms 65, 66soil inositol phosphate content 192

Analytical separation methods 24–29Animal feeds see Feeds, animalAnimal manures see Manures, animalAnimal nutrition 133–143

see also Dietary manipulationAnions 174, 225, 231–232appA-encoded phosphatase see Escherichia coli, AppA

phytaseAquaculture 122Aquatic systems 261–274

amounts of inositol phosphates 264–270phosphorus remobilization at sediment–water

interface 270–273physicochemistry of inositol phosphates 262–264sources of inositol phosphates 261–262see also Water bodies

Arabidopsispurple acid phosphatases 244seed phytic acid biosynthesis 114, 115–116transgenic phytase-expressing 102, 104–105, 246vacuolar adenosine triphosphatase 117

Arbuscular mycorrhizae 243, 248–250, 252Aspergillus expression systems 141Aspergillus fumigatus phytase 99, 101, 138, 140, 141, 222Aspergillus niger (formerly A. ficuum) phytases (mainly

PhyA) 64, 67, 80, 82in animal feeds 135–136, 173Apase6 104, 105applications 68, 102catalytic mechanisms 98, 99–100disulphide bridges 101expression systems 68, 100, 141glycosylation 100, 101, 142

Aspergillus niger (Continued )improving pH profile/catalytic properties

140–141PhyB 100properties 67–68, 69, 223proteolysis resistance 139soil interactions 223, 225, 226–227, 228, 229,

230–231transgenic plants overexpressing 138, 246, 247,

252–253Aspergillus oryzae phytase 222Aspergillus terreus phytase 69, 98Axenic culture 245

Bacillus spp. phytases 64–65, 66, 67, 68, 89applications 103catalytic mechanism 102–103degradation pathways 79, 81expression systems 68, 141–142inoculation of plants 245properties 69, 222regulation of synthesis 82, 83

Baculovirus expression system 142Barley

low phytate (lpa) varieties 113, 118–119, 126animal feeding studies 122, 138, 155, 156human feeding studies 124, 126

phytases 80, 87seeds 14, 15, 111

Barrel medic (Medicago truncatula) phytase 104–105, 246β-propellor phytases (BPP) 61, 71, 79, 102–103, 106Biofarming 102Bis-diphospho-myo-inositol tetrakisphosphate (IP8) 3,

114, 116Burkholderia spp. 70, 245

Calcareous soils 159, 160–161, 193, 211Calcite 211, 263Calcium

bioavailability, low-phytate crops 121, 122, 124,125

faecal excretion 135Calcium carbonate 212Calcium ions

effects on adsorption 216, 263inositol phosphate reactivity 171–172phytase activity and 82–83, 102, 103, 231

Calcium/phosphorus ratio, dietary 136, 137Calcium phytate 171, 172

adsorption in soil 211in culture media 62, 63precipitation in soils 213utilization by plants 247–248, 249

cAMP receptor protein (CRP) 82, 84Capillary electrophoresis 27–29

Carbohydrases 137Carbon

content, animal manures 159, 160, 161, 162,164

source, phytase synthesis and 82, 84–85starvation, phytase synthesis and 81, 82,

83, 84Carbon-14 (14C) labelling studies 50–53, 56–58, 199,

200Cations

adsorption in soils and 215–216complexation in soils 213inositol phosphate hydrolysis and 175inositol phosphate reactivity 171–173phytase activation/inhibition in soil 231

Cattail (Typha spp.) 79Cattle 70, 126, 173Caulobacter crescentus 88Cellulase 137–138Cereal grains 111–112Chelating agents 173–174, 232Chemical degradation 7, 8Chilton Conference on Inositol and Phosphoinositides

(1984) 1Citrate 63, 134, 136–137, 232Citrobacter braakii phytase 66, 67, 69, 139Clays

adsorption of phytases 223, 224, 228–229retention of organic phosphorus 207, 208sorption of inositol phosphates 212, 263

Clover, subterranean (Trifolium subterraneum) 246, 247,251, 252

Conformational inversion 8–10, 11Conformational isomers 8, 9Conformers 2Consensus-1 phytase 139, 141Consensus-7 phytase 141Copper chelate stability 173–174Cultivated soils 192, 193Culture media 62–63, 66–67Cyclic adenosine monophosphate (cAMP) 82, 841,2-Cyclohexanediamine tetraacetate (CDTA) 174,

175, 176, 177–178Cysteine phosphatases 61, 103–104Cysteine phytases (CPhy) 104, 106

Denaturing gradient gel electrophoresis (DGGE)71

Desorption, inositol phosphates 213–214, 271Diet, phosphorus composition of manures and 153,

154Dietary manipulation

environmental fate of manure phosphorus and161–163

future research needs 164–165overall benefits 164

280 Index

phosphorus content of manure 126, 133–134,135, 153–161

see also Feeds, animalDiethylene triaminepentaacetate (DTPA) 175, 176,

177–178Diphospho-myo-inositol pentakisphosphate (IP 7) 3,

114, 116Diphospho-myo-inositol tetrakisphosphate (PP-IP4) 3Drying, soil, phytase activity and 233Duckweed 262

Ecology, phytate-degrading microorganisms 68–71Ectomycorrhizal fungi 243, 248–250, 251–252EDTA see EthylenediaminetetraacetateElectrospray ionization (ESI) 23, 29–30Electrospray ionization–time of flight–mass

spectrometry (ESI–TOF–MS) 23–24, 29–38direct 31–34principles 29–30size-exclusion chromatography with 34–37, 38

Enterobacter 66, 89Enterobacter cloacae 78, 79, 80Enterobacteriaceae 90Environmental fate, manure phosphorus 161–163,

262–263Environmental issues 111–112, 133, 150–165Environmental samples

high-performance chromatography and massspectrometry 25, 34, 38

NMR spectroscopy 18–19Epimerization 2

microbial, in soils 49–50, 55–58reactions 199

Epimers 2Erosion, soil 161Escherichia coli

agp-encoded acid phosphatase 66, 83, 88AppA phytase 66, 67, 83–85, 88, 134–136

expression systems 141with improved thermostability 139improving pH profile/catalytic properties

140proteolysis resistance 139–140

expression systems 68, 102, 141phytases 64, 65–67, 78, 91

catalytic mechanism 98, 99degradation pathways 79, 80disulphide bridges 101expression systems 68, 100, 138, 141glycosylation 138, 139in vivo function 88–89, 90properties 69, 222regulation of synthesis 81, 82, 83–85

Ethylenediaminetetraacetate (EDTA)-exchangeable phytase-hydrolysable phosphorus

179–180

extraction methods 152–153, 190, 191ligand exchange studies 174, 175, 176, 177–178

Exchange spectroscopy (EXSY), random delay 10, 19Extraction methods

animal manures 152–153, 170–171aquatic systems 263, 266–269soil 41–42, 190

Feeds, animalpelleting 138phytase supplements see under Phytasesphytate content 111–112, 133using low-phytate crops see Low-phytate cropssee also Dietary manipulation

Fermentation technologies, phytase production 67Ferrihydrite 208, 209, 210, 213Fertilizers, phosphorus-based 102, 242

effects on soil inositol phosphates 193phytase-expressing transgenic plants 252

Fish, diets using low-phytate crops 122Flow-scintillation analysis, isotopic tracers in soils

54–55, 56Food, applications of phytases 102Forest soils 192Formic acid 84, 136–137Fractionation, phosphorus in animal manures 152,

170Fragmentation, mass spectral ion

inositol phosphates 32–34, 35, 37, 38inositol stereoisomers and products 43–46

Freezing, phytase denaturation 233Fungal phytases 65, 67, 82, 99–100, 222

applications 102consensus constructs 139, 141disulphide bridges 101glycosylation 100, 101, 142transgenic plants overexpressing 246–247

Gas chromatography 24Gas–liquid chromatography 24Gel chromatography 25Germination, phytate degradation 87Gibberellic acid 87Globoids 111, 116–117Glossary of terms 2–5Glucose, effects on phytase synthesis 82Glucose-1-phosphatase 78, 83Glycosylation

Escherichia coli phytase 138, 139fungal phytases 100, 101, 142phytases in soil 230

Goethite (Fe(OOH))adsorption to 209, 210, 211, 214–215, 263desorption from 213, 271

Grassland soils 192, 195

Index 281

Hebeloma cylindrosporum phytase 225, 228Hedley fractionation method 152, 170Heteronuclear multiple quantum correlation plus two-

dimensional total correlation spectroscopy(HMQC-TOCSY) 15–17, 20

High-performance liquid chromatography (HPLC)24, 54

reversed-phase (RP-HPLC) 23, 25, 26, 27High-performance size-exclusion chromatography

(HP-SEC) 23, 34–37mass fragmentometry approach 37, 38principle and technique 34–36selected ion monitoring (SIM) approach 36–37,

38Histidine acid phosphatases (HAP) 61, 98–102

disulphide bridges 101glycosylation 100phytate degradation pathway 79substrate specificity site 99–100

Histidine acid phytases (HAPhy) 99–102, 106Human nutrition 112

low-phytate crops 122–126, 127–128phytase supplements 135

Humic materials 191, 212–213, 223–224, 228Hydrogen-bonding interactions 18Hypobromite oxidation (alkaline bromination) 190,

191, 196–197, 264, 265, 267, 268

Identification of inositol phosphatesin aquatic systems 266–269by mass spectrometry see Mass spectrometryby NMR spectroscopy 7–20

in environmental samples 18–19in impure samples 15–17in a mixture without separation 12–14in plant seeds 14purified compounds 10–12

in soils 18–19, 46, 48, 49, 186, 190, 191Inductively coupled plasma (ICP) mass spectrometry

(ICP–MS) 23, 27Infrared spectroscopy 46, 47Inositol 2allo-Inositol 3, 198, 199chiro-Inositol

infrared spectroscopy 46, 47proton NMR spectroscopy 46, 48in soil 57

D-chiro-(+)-Inositol 3, 194mass spectrometry 43, 44origins 49–50, 55, 57, 58, 198, 199in soils 49–50, 194

identification 46–49isotopic studies 50, 53, 54

L-chiro-(−)-Inositol 3, 198, 199cis-Inositol 3, 198, 199epi-Inositol 3, 198, 199

muco-Inositol 3identification in soils 47origins 198, 199

myo-Inositol 2, 3, 194Agranoff’s turtle 4infrared spectroscopy 46, 47mass spectrometry 43–46NMR spectroscopy 16–17, 46, 48origins 198, 199in soil 50, 194

carbon and phosphorus pathways 55identification methods 43–46, 47, 48isotopic studies 50–53, 54, 56–58microbial epimerization 49–50, 55–58

neo-Inositol 3, 194origins 57, 198, 199in soils 49, 50

scyllo-Inositol 3, 5, 194origins 57, 198, 199in soils 49, 50, 194

myo-Inositol bisphosphate 3, 32, 36, 37D-chiro-(+)-Inositol hexakisphosphate 195, 196, 197muco-Inositol hexakisphosphate 49, 195, 198, 199myo-Inositol hexakisphosphate (InsP6) 2, 3, 4, 97

see also phytic acidin animal manures 150, 151–152, 153,

154, 170dietary manipulation 153–158environmental fate 162, 163ligand effects 174–180phosphorus solubility in soil and

159–161reactivity with polyvalent cations 173

in aquatic systems 261–274biosynthesis in seeds 114–116cation complexes 171–173

ligand exchange 174–178pH effects 172–173in ruminant excreta 173in soils 213, 214stability calculations 174

conformational inversion 8–9, 10, 11identification by NMR spectroscopy 14, 16–17,

190, 191identification of hydrolysis products 13–14mass spectrometry 32–33, 34phosphorylases see PhytasespKa values 18, 214, 215salts see Phytatesize-exclusion chromatography and mass

spectrometry 36–37, 38in soils 187–188, 189, 193, 194

adsorption 209, 210–211, 212, 214–217complexation 213, 214desorption 213, 214isotopic studies 50–53origins 199

282 Index

neo-Inositol hexakisphosphate 195, 196, 197scyllo-Inositol hexakisphosphate

NMR spectroscopy 18–19, 190, 191origins 195, 199potential function 201in soils 194–197

Inositol hexakisphosphatesorigins 195–199in soils 187, 188–189, 196–197

myo-Inositol kinase 114, 115myo-Inositol monophosphate 3

mass spectrometry 32, 33, 34NMR spectroscopy 12, 16size-exclusion chromatography and mass

spectrometry 36–37D-myo-Inositol 3-monophosphate synthase (MIPS)

114–115, 120myo-Inositol pentakisphosphate 3

biosynthesis in seeds 114, 116conformational inversion 10, 11mass spectrometry 32NMR spectroscopy 14

Inositol pentakisphosphates, in soil 187–190Inositol phosphates 2, 3

in animal manures 150–165, 195–199in animal nutrition 133–143in aquatic systems 261–274identification see Identification of inositol

phosphatesnomenclature 1–5plant utilization 242–254separation and detection by mass spectrometry

23–38in soil see under Soils

D-chiro-(+)-Inositol phosphates 195, 198myo-Inositol phosphates 3

in aquatic systems 261–262in low phytic acid seeds 112–113metabolic pathways in seeds 113–116origins 198in soils 194

neo-Inositol phosphates 194, 195, 198scyllo-Inositol phosphates

origins 195, 198in soils 194–195

myo-Inositol polyphosphate 2-kinase 114, 116Inositol stereoisomers (and phosphorylated

derivatives) 3, 5in aquatic systems 262, 269in soils 41–58, 193–201

myo-Inositol tetrakisphosphate 3conformational inversion 10, 11mass spectrometry 32NMR spectroscopy 14

myo-Inositol trisphosphate 3, 8, 18, 32myo-Inositol 1,3,4-trisphosphate 5/6-kinase 114, 115,

116

myo-Inositol 1,4,5-trisphosphate 3/6-kinase 114, 115,116

Inosose, mass spectrometry 43–46DL-epi-Inosose, mass spectrometry 43, 44International Union of Pure and Applied

Chemistry (IUPAC) and InternationalUnion of Biochemistry (IUB) 1, 4–5, 7, 78

Ion chromatography 26–27, 28Ion-exchange chromatography 24, 26

radionuclide tracers in soils 54, 56Ion-pairing reversed-phase high-performance liquid

chromatography (HP-ion pair-RPLC) seeunder High performance liquid chromatography23, 25, 27

Iron bioavailabilitydietary phytase supplements 135low-phytate crops 123–124

Iron hydroxide, addition to soil 180, 181Iron oxides

adsorption to 208–211, 263desorption from 213–214, 271

Iron(II) phytate, in aquatic systems 271Iron(III) ions

chelate stability 173–174inositol phosphate hydrolysis and 177–178inositol phosphate reactivity 171–172, 172sorption in aquatic systems and 263, 270, 271

Iron(III) phytate, in soils 50–53, 192, 213Isoelectric points, phytases 222, 225, 227Isomers

conformational 8, 9positional 4–5

Isotopic tracer studies, in soils 50–58, 199, 200, 248,252–253

Kaolinite 208, 209–210, 212, 228–229Klebsiella spp. phytases 66, 69, 89

purification and characterization 64, 65regulation of synthesis 82

Lactic acid 136–137, 138Lactic acid bacteria (Lactobacillus spp.) 66, 69, 88,

138, 141–142Legumes 111, 112Ligands

based fractionation assay 178–180exchange processes 174–178, 272–273sources 174stability of cation complexes 173–174

Light radiation, phytase degradation 234Lily (Lilium longiflorum), phytate-degrading enzymes

79, 81, 87, 103Low-phytate crops 112–113, 117–128

animal feeding 120–122, 155, 156

Index 283

Low-phytate crops (Continued )combined with phytase supplements 158,

159, 163environmental fate of manure phosphorus

161–162faecal phosphorus excretion 121, 122, 135

human nutrition 122–126seed phosphorus and ruminant nutrition and

126–127low phytic acid (lpa) genotypes 112–113, 114, 115–116,

117–119Lucerne 102, 252Lupin (Lupinus spp.)

phytases 79, 80, 244, 246phytate-degrading microorganisms 70utilization of soil phytates 247–248

Magnesium, in low-phytate rice 125–126Maize

low phytate (lpa) varieties 112, 113, 117, 118, 119animal feeding studies 120–121, 122, 155human nutrition studies 123–125phytase supplements with 138seed phytic acid biosynthesis 114, 115–116

phytases 87, 98, 246seeds 111, 112utilization of soil phytate 248

Manures, animal 150–165carbon/phosphorus ratios 159, 160, 161, 162,

164environmental issues 111–112, 133, 150–165inositol phosphates 150–165, 195–199inositol phosphates reaching aquatic systems

262–263nitrogen/phosphorus ratios 151, 153phosphorus 150–165

analytical methods 151–153, 170–171composition 151–153, 154dietary manipulation 126, 133–134, 135,

153–161environmental fate and dietary manipulations

161–163phosphorus solubility in soil and 158–161storage effects 158, 164temporal changes in biological availability

180, 181solubility and release of inositol phosphates

169–181analytical methods 169–171, 178–180characterizing relative stability 173–180ligand exchange studies 174–178reactivity with polyvalent cations 171–173

Mass fragmentometry 37, 38Mass spectrometry (MS) 23–24, 29–38

capillary electrophoresis with 27direct 31–34

electron-impact 43–46, 49ion-pairing reversed-phase HPLC with 25, 27radionuclide tracers in soils 54size-exclusion chromatography with 34–37

Metal ionscomplexation of inositol phosphates 70, 71,

173–174, 213, 263phytase inhibition 231

Microorganismsdegradation of soil phytases 230–231phosphorylated inositol stereoisomers 199, 201phytases see Phytases, microbialphytate-utilizing see Phytate-degrading

microorganismssoil, phosphorus utilization by plants and 243,

248, 251–252synthesis of inositol phosphates 195, 199–201

Mineral nutrition, human 112, 123–126Monogastric animals 70, 111–112

dietary manipulation of manure phosphorus153–161

dietary phosphate supplements 153dietary phytase supplements 156–158diets using low-phytate crops 120–122, 155, 156environmental fate of manure phosphorus

161–163, 262–263phosphorus composition of manures 153, 154,

170Montmorillonite 212, 224, 228–229Mung bean phytase 79, 80, 87Mycorrhizal fungi 243, 244, 248–250, 251–252

Nitrate, repression of phytase synthesis 85Nomenclature 1–5Non-ruminant animals see Monogastric animalsNuclear magnetic resonance (NMR) spectroscopy

7–20acid dissociation constants 17–18animal manures 152–153, 154applications 9aquatic system samples 269conformational analysis 8–10environmental samples 18–19experimental details 19–20intramolecular hydrogen bonding 18plant root studies 251protonation sequences at microscopic level

17–18radionuclide tracers in soils 54–55soil samples 18–19, 46, 48, 49, 186, 190, 191solid-state 19structural determinations 10–17TOCSY technique 12–14, 15

Nutrient statusregulation of phytase synthesis 88–89soil inositol phosphates and 193, 195

284 Index

Organic matter, soil 212–213, 223–224, 228Oxalic acid 175, 176, 177, 178, 232

Pantoea agglomerans 79, 80Paper chromatography 24–25, 41–42Paramecium 66, 79, 80, 201Pathogenic infections 89–90, 103Pelleting, animal feed 138Penicillium simplicissimum 69Peniophora lycii phytase 68, 140

interactions in soil 225, 227, 229, 230–231properties 69, 222

pHinositol phosphate complexation and 213, 214inositol phosphate hydrolysis and 172–173optima of phytases 140–141, 229, 232phosphorus mobilization in aquatic systems

272–273phytase synthesis and 82soil

abiotic processes and 214–215, 225, 226inositol phosphate content and 192–193phytase activity and 232

PHO regulon 67, 83, 85–86Phosphatases

hydrolysis of soil organic phosphorus 243–244see also Phytases

Phosphate, inorganicanimal feed supplements 111, 153in animal manures 152, 173–180fertilizers see Fertilizers, phosphorus-basedinhibition of phytases in soil 231–232ligand-exchangeable 179microbial assimilation 88regulation of phytate-degrading activity 87seed 111, 112, 113, 117, 119uptake by plants 243see also Phosphorus

Phosphatidylinositol (PtdIns) phosphates, in seeds 114,115, 116, 117

Phosphoinositides 1Phospholipase C 89–90Phosphomonoesterases 78

see also phosphatasesPhosphorus (P)

in animal manures see under Manures, animalbioavailability

animal feedstuffs 133–134low-phytate crops 121, 122

dietary manipulation strategies see Dietarymanipulation

environmental issues 111–112, 133, 150–165faecal excretion

dietary phytase supplements and 135low-phytate crops 121, 122, 135manipulation strategies 133–134, 153–158

fertilizers see Fertilizers, phosphorus-basedlimitation/deficiency

effects on plants 242, 243–245inositol phosphate levels and 70, 193, 195phytase synthesis and 66, 81, 83, 84,

85–86, 89phytase-hydrolysable (PHP) see Phytase-

hydrolysable phosphorusseed 111, 112

crops with reduced total 126–127in low-phytate crops 113, 119–120non-ruminant nutrition studies 120–122

soilaccumulation 151run-off to water bodies 163, 164solubility, after manure application

158–161, 164utilization by plants 242–254

see also Inositol phosphates; Phosphate, inorganic

Phosphorus-32 (32P) tracer studies 55, 58, 248Phosphorus-33 (33P) tracer studies 56, 58, 252–253Phthalic acid 175–176, 177, 178Phytase-hydrolysable phosphorus (PHP)

EDTA-exchangeable 179–180ligand-based assay 178–180in soils 193, 251, 252

Phytases (phytate-degrading enzymes) 4, 61, 78–91animal feed supplementation 68, 102, 103,

133–143activity in stored manures 136, 158augmentation strategies 137–138combined with low-phytate diets 158, 159,

163constraints 138determinants of efficacy 136–137enhancing proteolysis resistance 139–140environmental fate of manure phosphorus

162–163impact on manure phosphorus 135,

156–158improving pH profile/catalytic properties

140–141nutritional impacts 134–135production systems 141–142site of activity in animals 135–136thermostability 138–139

applications 102, 103attributes and catalytic mechanisms 97–105classification 61, 78–79, 97–98, 105, 106in vivo function 87–90ligand exchange effects 174–180microbial 78

characterization of activity 63–65degradation pathways 79, 80, 81expression and production 65–67, 68in vivo function 88–90

Index 285

Phytases (Continued )intra- and extracellular 65, 66, 89–90ions and metabolites activating/inhibiting

231–232properties 67–68, 69, 222purification 63, 64regulation of synthesis 79, 81–83sources 65, 66

nomenclature 105, 106phosphorylated stereoisomer hydrolysis 201plant 78, 79

constitutive 86–87degradation pathways 79, 80germination-inducible 86, 87in vivo function 88ions and metabolites activating/inhibiting

231–232regulation of synthesis 86–87role in uptake of soil organic phosphorus

243–244, 245plant roots 88, 244, 245–247production systems 65–66, 67, 138, 141–142regulation of synthesis 79–87in soil 221–235

denaturation 232–233factors affecting activity 222–223ions and metabolites inhibiting/activating

231–232microbial and proteolytic degradation

230–231solid phase interactions 223–230

in soil–plant root environment 243–244transgenic animals 221, 263transgenic plants see Transgenic plants, phytase-

expressing3-Phytases 78, 796-Phytases 78, 79Phytate 2, 4

in animal feeds 111–112, 133in animal manures 153in human diet 112phosphorus content 97seed 111–128utilization by plants 245, 247–248, 249,

250see also myo-Inositol hexakisphosphate

Phytate-degrading enzymes 4, 78see also Phytases

Phytate-degrading microorganisms 61–72, 250–251in aquatic systems 271–272assessment 61–65ecology 68–71inoculation of plants with 245isolation case study 63screening for 62–63sources 65, 66see also Phytases

Phytic acid 4, 111see also myo-inositol hexakisphosphate and phytatebiosynthesis in seeds 113–116

Phytins 2, 4, 111Phytoplankton 262Pichia pastoris expression systems 68, 140, 141Pigs see SwinePinitol 198, 199Pinus spp. 244, 250, 251Pisolithus tinctorius 225pKa values 17–18, 214, 215Plants

aquatic 262in axenic culture 245mycorrhizal associations 243phytate-degrading enzymes see under Phytasessynthesis of inositol stereoisomers 199–201transgenic see Transgenic plantsutilization of inositol phosphates 242–254see also Rhizosphere; Roots; Seeds

Pollen, phytate-degrading enzymes 79, 86–87Polymerase chain reaction (PCR) 71Positional isomers 4–5Potassium, in low-phytate rice 125–126Poultry 70, 97

dietary manipulation of manure phosphorus155, 156–158, 159

dietary phytase supplements 134–135, 136–137,156–158, 162–163

environmental fate of manure phosphorus162–163, 262–263

low-phytate crop-based diets 120–121, 155, 156

low-phytate grains plus phytase supplements158, 159

phosphorus composition of manures 153, 154Precipitation, cation complexes in soils 214, 246Preparative separations 24, 41–42Proteases 137–138Protein storage vacuoles (PSVs) 111, 116–117Protein tyrosine phosphatase (PTP) 104Proteolysis, phytases 139–140, 230–231Protonation sequence, at microscopic level 17–18Protozoa 201

see also Tetrahymena vorax and ParmeciumPseudomonas spp.

phosphate utilization 89phytases 69, 80, 82, 245phytate degradation 62, 63, 66

Pteris vittata 202Purple acid phosphatases (PAPs) 61, 104–105,

244, 246Purple acid phytases (PAPhy) 104–105, 106Pyrophosphatases 103

Quebrachitol, mass spectrometry 43, 44, 45, 46

286 Index

Radioisotope labelling studies 50–54, 199, 200, 248,252–253

Raoultella terrigena phytase 79, 81, 82Rhizopus oligosporus 66Rhizosphere

microorganisms 70, 88phosphate uptake by plants 243, 248, 251,

252phytases 224–225, 232, 244

Ricebran 82low phytic acid (lpa) 113, 125–126pathogens 90, 103phytases 80, 104seeds 111

Rivulariaceae 88RNA polymerase, σs subunit 83–84Roots

phosphorus uptake from soils 243, 244–245phytate-degrading enzymes 88, 244, 245–247phytate-degrading microorganisms 70see also Rhizosphere

Rumen bacteria 89, 103–104Ruminants 70, 112

low seed total phosphorus crops 126–127myo-inositol hexakisphosphate in excreta 173phosphorus composition of manures 153, 154,

170Rye phytase 80, 87Ryegrass Lolium perenne L. 248, 251–252

Saccharomyces carlsbergensis 201Saccharomyces cerevisiae

expression systems 68, 139–140, 141phytases 69, 80, 91, 222

in vivo function 88production and expression 65–66, 67regulation of synthesis 85–86

Salinity, changes in 273Salmonella dublin 90Schwanniomyces castellii 69, 82Second messengers 89–90Sediments

amounts of inositol phosphates 266–270phosphorus remobilization mechanisms

270–273physical and biological perturbation 273sources of inositol phosphates 261–262

Seedsfield emergence 120inositol phosphates 112–113NMR spectroscopy 14, 15phosphorus see Phosphorus (P), seedphytate 111–128

deposition in globoids 116–117lpa genotypes 112–113

metabolic pathways, genes and mutants113–116

see also Low-phytate cropsphytate-degrading enzymes 79, 86–87, 88

Selected ion monitoring (SIM) 33, 36–37, 38Selenomonas ruminatum phytase 66, 82, 89, 103–104Separation methods

analytical 23, 24–29preparative 24, 41–42radionuclide tracers in soils 54

Sewage sludge 195Shewanella oneidensis 71Size-exclusion chromatography see High-performance

size-exclusion chromatographySodium hydroxide (NaOH) extractions 152–153, 170,

172, 190, 191Sodium ions, inositol phosphate reactivity 171,Sodium phytate 171

in culture media 62, 63utilization by plants 245, 247–248

Soilsinositol phosphates 186–202

abiotic reactions 207–217amounts 186–190extraction and preparative chromatography

41–42factors controlling amounts 190–193NMR spectroscopy 18–19, 46, 48, 49,

186, 190, 191reaching aquatic systems 262

inositol stereoisomers (and phosphorylated deriv-atives) 41–58, 193–201

isotopic studies 50–58methodologies for characterizing 41–49origins 49–50, 195–201potential function 201significance 49–50

iron hydroxide addition to manured 180, 181ligand-based phytase-hydrolysable phosphorus

assay 178–180phosphorus accumulation 151phosphorus solubility in manure-treated

158–161, 163, 164phosphorus uptake by plants 243–245phytases in see under Phytasesphytate-degrading microorganisms 63, 65, 66,

70, 71, 250–251phytate utilization by plants 245, 247–248,

249Sorption see AdsorptionSoybean

low phytate varieties 113, 119, 120, 121phytase (GmPhy) 79, 104, 105, 222, 246seeds 111transgenic phytase overexpressing 138

Spirodela polyrhiza 90Stationary phase response 81, 83, 85, 89

Index 287

Stereoisomers 5see also Inositol stereoisomers

Stress tolerance, low-phytate crops 118Suillis phosphatases 225Surface properties, effects of sorption 216–217Swine

dietary manipulation of manure phosphorus155, 156, 158, 159

dietary phytase supplements 134–138, 156,162–163

diets using low-phytate crops 122, 135, 155,156, 162–163

environmental fate of manure phosphorus162–163, 262–263

Escherichia coli phytase expression 99low-phytate grains plus phytase supplements

158, 159phosphorus composition of manures 153,

154

Temperature effectsphytase stability 138, 232–233phytase synthesis 82

Terminal-restriction fragment length polymorphisms(T-RFLP) 71

Terminology 1–5Tetrahymena vorax 198, 201Thermostable phytases 138–139Time of flight (TOF) mass spectrometry 23–24, 30,

31–34Tobacco (Nicotiana tabacum) 103, 246, 248, 249,

252TOCSY see Two-dimensional total correlation

spectroscopyTransgenic animals, phytase-expressing 221, 263Transgenic plants, phytase-expressing 102, 103,

104–105, 221, 246–247in animal feeds 138, 141growth and phosphorus nutrition 248, 252–253

Trifolium subterraneum see Clover, subterranean

Tritium (3H) tracer studies, in soil 56–58Turtle (structure) 4, 5Two-dimensional total correlation spectroscopy

(TOCSY) 12–14, 15heteronuclear multiple quantum correlation

(HMQC) 15–17, 20technique 19–20

Type III secretion systems 89–90

Ultraviolet (UV) absorbance detection 54

Vacuolar adenosine triphosphatase (V-ATPase) 116,117

Vitamin D derivatives 134, 136

Water bodiesamounts of inositol phosphates 264, 265, 266transport of manure phosphorus to 151,

158–161, 163, 164, 262–263see also Aquatic systems

Water content of soil, phytase activity and 233Wetland soils 192, 269–270Wheat

bran 67, 82low phytic acid (lpa) genotypes 113, 126phytase 79, 87, 245–246seeds 111utilization of phytates 248

Wolffiella floridana 262

Xanthomonas oryzae 90, 103

Yields, low-phytate crops 118–119

Zinc 124–125, 173–174

288 Index

Inositol Phosphates: Linking Agriculture and the Environment

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